Nitrated lipids and methods of making and using thereof

ABSTRACT

Described herein are nitrated lipids and methods of making and using the nitrated lipids.

CROSS REFERENCE TO RELATED APPLICATIONS

This application is a continuation of U.S. patent application Ser. No.12/797,460, filed Jun. 9, 2010, which claims priority to divisional ofU.S. patent application Ser. No. 11/568,377, filed Oct. 26, 2006, whichclaims priority to U.S. Patent Application Ser. No. 60/566,005, filedApr. 28, 2004, the entire disclosures of which are hereby incorporatedby reference in their entireties for all purposes.

ACKNOWLEDGEMENTS

This invention was made with government support under Grant NumbersRO1HL58115 and RO1HL64937 awarded by the National Institutes of Health.The government has certain rights in the invention.

BACKGROUND

Nitric oxide (.NO) is an endogenously generated, lipophilic signalingmolecule that maintains vascular homeostasis via stimulation of solubleguanylate cyclase (1). In addition to mediating vascular relaxation, .NOpotently modulates oxygen radical reactions, inflammatory cell function,post-translational protein modification and regulation of geneexpression (2-5). There are multiple pathways whereby .NO-derivedspecies can mediate the oxidation and nitration of biomolecules such asunsaturated fatty acids. Nitric oxide reacts at diffusion-limited rateswith superoxide (O₂.⁻, k=1.9×10¹⁰ M⁻¹ sec⁻¹) to yield peroxynitrite(ONOO⁻) and its conjugate acid, peroxynitritrous acid (ONOOH), thelatter of which undergoes homolytic scission to nitrogen dioxide (.NO₂)and hydroxyl radical (.OH) (2, 6). Also, biological conditions favor thereaction of ONOO⁻ with CO₂, yielding nitrosoperoxycarbonate (ONOOCO₂ ⁻;k=3×10⁴ M⁻¹ sec⁻¹), which rapidly yields .NO₂ and carbonate (.CO₃ ⁻)radicals via homolysis, or rearrangement to NO₃ ⁻ and CO₂ (7). Duringinflammation, neutrophil myeloperoxidase and heme proteins such asmyoglobin and cytochrome c catalyze H₂O₂-dependent oxidation of nitrite(NO₂ ⁻) to .NO₂, resulting in biomolecule oxidation and nitration thatis influenced by the spatial distribution of catalytic heme proteins(8-11). Finally, even though the rate of reaction of .NO with O₂ isslow, (k=2×10⁶ M⁻² sec⁻¹) the small molecular radius, uncharged natureand lipophilicity of .NO and O₂ facilitate their diffusion andconcentration in membranes and lipoproteins up to 20-fold (12-14). This“molecular lens” effect induced by .NO and O₂ solvation in hydrophobiccell compartments accelerates the reaction of .NO with O₂ to yield N₂O₃and N₂O₄. As a result of these various reactions, a rich spectrum ofprimary and secondary reactions yield products capable of concertedoxidation, nitrosation and nitration of target molecules.

Multiple mechanisms can account for the nitration of fatty acids by .NO₂(15-20). During both basal cell signaling and tissue inflammatoryconditions, .NO₂ generated by the aforementioned reactions can reactwith membrane and lipoprotein lipids. Environmental sources also yield.NO₂ as a product of photochemical air pollution and tobacco smoke. Inboth in vivo and in vitro systems, .NO₂ has been shown to initiateauto-oxidation of polyunsaturated fatty acids via hydrogen abstractionfrom the bis-allylic carbon to form nitrous acid and aresonance-stabilized allylic radical (21). Depending on the radicalenvironment, the lipid radical species can react with molecular oxygento form a peroxyl radical. During inflammation or ischemia, when O₂levels are lower, lipid radicals can react to an even greater extentwith .NO₂ to generate multiple nitration products including singlynitrated, nitrohydroxy- and dinitro-fatty acid adducts (18, 19, 21).These products can be generated via either hydrogen abstraction ordirect addition of .NO₂ across the double bond. Hydrogen abstractioncauses a rearrangement of the double bonds to form a conjugated diene;however, the addition of .NO₂ maintains a methylene-interrupted dieneconfiguration to yield singly nitrated polyunsaturated fatty acids (18).This arrangement is similar to nitration products generated by thenitronium ion (NO₂ ⁺), which can be produced by ONOO⁻ reaction with hemeproteins or via secondary products of CO₂ reaction with ONOO⁻ (20).

Reaction of polyunsaturated fatty acids with acidified nitrite (HNO₂)generates a complex mixture of products similar to those formed bydirect reaction with .NO₂, including the formation of singly nitratedproducts that maintain the bis-allylic bond arrangement (18, 19). Theacidification of NO₂ ⁻ creates a labile species, HNO₂, which is inequilibrium with secondary products, including N₂O₃, .NO and .NO₂, allof which can participate in nitration reactions. The relevance of thispathway as a mechanism of fatty acid nitration is exemplified byphysiological and pathological conditions wherein NO₂ ⁻ is exposed tolow pH (e.g., <pH 4.0). This may conceivably occur in the gastriccompartment, following endosomal or phagolysosomal acidification or intissues following-post ischemic reperfusion.

Nitrated linoleic acid (LNO₂) displays robust cell signaling activitiesthat (at present) are anti-inflammatory in nature (20, 22-25). SyntheticLNO₂ inhibits human platelet function via cAMP-dependent mechanisms (26)and inhibits neutrophil O₂.⁻ generation, calcium influx, elastaserelease, CD11b expression and degranulation via non-cAMP,non-cGMP-dependent mechanisms (27). LNO₂ also induces vessel relaxationin part via cGMP-dependent mechanisms (22, 28). In aggregate, thesedata, derived from a synthetic fatty acid adduct, infer that LNO₂species represent a novel class of lipid-derived signaling mediators. Todate, a gap in the clinical detection and structural characterization ofnitrated fatty acids has limited defining LNO₂ derivatives asbiologically-relevant lipid signaling mediators that converge .NO andoxygenated lipid signaling pathways.

Therefore, it would be advantageous to produce nitrated lipids insubstantially pure form so that their cell signaling activities can becharacterized and their purified derivatives can be used to treatvarious diseases. Described herein are nitrated lipids and methods forproducing nitrated lipids in pure form. Also described herein aremethods for using the nitrated lipids to treat various diseases.

SUMMARY

Described herein are nitrated lipids and methods of making and using thenitrated lipids. The advantages of the invention will be set forth inpart in the description which follows, and in part will be obvious fromthe description, or may be learned by practice of the aspects describedbelow. The advantages described below will be realized and attained bymeans of the elements and combinations particularly pointed out in theappended claims. It is to be understood that both the foregoing generaldescription and the following detailed description are exemplary andexplanatory only and are not restrictive.

BRIEF DESCRIPTION OF THE DRAWINGS

The accompanying drawings, which are incorporated in and constitute apart of this specification, illustrate several embodiments of theinvention and together with the description, serve to explain theprinciples of the invention.

FIG. 1 shows a reaction scheme for producing nitrated lipids.

FIG. 2 shows the separation of C-10 and C-12 nitrated linoleic acid froma crude reaction mixture by thin layer chromatography.

FIG. 3 shows the extinction coefficient (A) and ultraviolet lightabsorption spectrum (B) of nitrated linoleic acid.

FIG. 4 shows the characterization of nitrated linoleic acid by GC massspectrometry.

FIG. 5 shows the HPLC resolution of individual positional isomers ofnitro derivatives of linoleic acid present in a) synthetic preparations,b) red cells and c) plasma; and in concert with this exemplification ofpositional isomer resolution is the structural characterization ofindividual nitrated linoleic acid positional isomers by electrosprayionization triple quadrupole mass spectrometry.

FIG. 6 shows the standard curve used for quantitative analysis of redblood cell and plasma nitrated linoleic acid content.

FIG. 7 shows that LNO₂ is a potent PPAR ligand. (A) CV-1 cells,transiently co-transfected with different nuclear receptor ligandbinding domains fused to the Gal4 DNA binding domain and the luciferasereporter gene under the control of four Gal4 DNA binding elements, wereincubated with vehicle (methanol) or LNO₂ (3 μM, 2 hr, n=4). (A, inset)Dose-response of LNO₂-dependent PPARγ ligand binding domain activation(n=4). (B) Dose-response of LNO₂-dependent PPARγ, α and δ activation(n=4). The luciferase reporter gene was under the control of three PPARresponse elements (C) Response of CV-1 cells transfected with PPARγ anda luciferase reporter construct under the control of PPRE followingexposure to LNO₂ and other reported PPARγ ligands (1 and 3 μM each ofciglitazone, 15-deoxy-Δ^(12,14)-PGJ₂,1-palmitoyl-2-hydroxy-sn-glycero-3-phosphocholine (LPA 16:0),1-oleoyl-2-hydroxy-sn-glycero-3-phosphocholine (LPA 18:1),1-O-hexadecyl-2-azelaoyl-sn-glycero-3-phosphocholine (AzPC),1-palmitoyl-2-azelaoyl-sn-glycero-3-phosphocholine (azPC ester),Δ^(9,11)-conjugated linoleic acid (CLA-1) and Δ^(10,12)-conjugatedlinoleic acid (CLA-2), with (n=3 to 5). “Vector” indicates empty vector(C, inset) Using the same reporter construct, the dose response of PPARγactivation by LNO₂, rosiglitazone, 15-deoxy-Δ^(12,14)-PGJ₂ and linoleicacid was measured (n=3). All values are expressed as mean±SD. (*)represents significantly different (P<0.05) from vehicle control usingStudent's t test. All experiments were repeated at least three times.

FIG. 8 shows the characterization of the PPARγ ligand activity of LNO₂.(A) Using CV-1 cells cotransfected with PPARγ and PPRE-controlledluciferase expression plasmids, the activation of PPARγ by LNO₂ wasevaluated in the absence or presence of PPARγ-specific antagonist GW9662added 1 hr prior to LNO₂ addition or upon co-addition of the RXRreceptor co-activating ligand 9-cis-retinoic acid (n=3). PPARγactivation by LNO₂ was inhibited in a dose-dependent manner by GW9662and was enhanced in the presence of the coactivator 9-cis-retinoic acid.(B) The action of LNO₂ as a PPARγ ligand was compared with LNO₂-deriveddecay products. Effective LNO₂ concentrations after selected decayperiods were measured by LC-MS with electrospray ionization using[¹³C]LNO₂ as internal standard (9). PPARγ activation was assessed viaPPRE reporter analysis in CV-1 cells. (n=3) (c) Potential PPARγ ligandactivity of LNO₂ decay products was measured via PPRE reporter analysis(n=4) (D) Competition of LNO₂, linoleate and unlabeled Rosiglitazone forPPARγ-bound [³H] Rosiglitazone. For (A-C), all values are expressed asmean±SD. (*) represents significantly different (P<0.05) from vehiclecontrol, and (#) represents significantly different from LNO₂ alone,using Student's t test. All experiments were repeated at least threetimes.

FIG. 9 shows that LNO₂ induces CD36 expression in macrophages andadipogenesis of 3T3-L1 preadipocytes. (A) Mouse RAW264.7 macrophages at˜90% confluence were cultured in DMEM with 1% FBS for 16 hours and thentreated with various stimuli for 16 hours as indicated. ThePPARγ-specific antagonist GW9662 was added 1 h prior to the treatment.The cell lysate was immunoblotted with anti-CD36 and anti-β-actinantibodies. (B,C) Two days after reaching confluence, 3T3-L1preadipocytes were cultured for 14 days and stained using Oil red O aspreviously (18) (B) or treated with various stimuli as indicated and thecell lysate was immunoblotted with anti-PPARγ, anti-aP2 and anti-β-actinantibodies (C). (D) LNO₂ increases [³H]-2-deoxy-D-glucose uptake in3T3-L1 adipocytes. Left panel: The dose-dependent effects of LNO2 on[³H]-2-deoxy-D-glucose uptake in 3T3-L1 adipocytes. Right panel:PPARγ-specific antagonist GW9662 was added 1 h before the treatment.[³H]-2-deoxy-D-glucose uptake assay was performed as described insupplemental methods. All experiments were repeated at least threetimes. Values are expressed as mean±SD (n=6). Statistical analysis wasdone by using Student's t test (*p<0.05 vs vehicle control; ^(#)p<0.05vs GW9662 untreated groups). Vehicle (Veh); Rosiglitazone (Rosi);15-deoxy-PGJ₂ (15-d-PGJ₂); linoleic acid (LA).

FIG. 10 shows nitrated oleic acid (OA-NO₂). Two regioisomers of OA-NO₂were synthesized by nitrosenylation of oleic acid and purified asdescribed in Experimental Procedures, generating 9- and10-nitro-9-cis-octadecenoic acids.

FIG. 11 shows nitrated fatty acid species in plasma and urine. Potentialnitroalkene products were evaluated in plasma and urine. Fatty acidswere extracted from clinical samples and analyzed by ESI-MS/MS asdescribed in Methods. Nitrated fatty acid adducts (—NO₂) and theirnitrohydroxy counterparts (L(OH)—NO₂) were detected using the multiplereaction monitoring (MRM) scan mode (Table 3) and are presented as baseto peak HPLC elution profiles with maximum ion intensity given on theleft axis. Six fatty acids were monitored: oleic acid (18:1), linoleicacid (18:2), linolenic acid (18:3), arachidonic acid (20:4)eicosapentaenoic acid (20:5) and docosahexaenoic acid (22:6). In plasmaand urine, all of the nitrated fatty acids and their Michael additionproducts with H₂O (nitrohydroxy adducts) appear in the HPLC elutionprofiles.

FIG. 12 shows ¹H and ¹³C NMR spectrometry of synthetic nitro-oleate(OA-NO₂). Proton (A) and ¹³C (B) NMR spectrometry confirmed thestructure of synthetic OA-NO₂. Identified protons and carbons areindicated for each regioisomer; downfield shifts are presented in ppm.¹³C NMR spectrometry indicates that synthetic OA-NO₂ is a mixture of tworegioisomers, with most carbon peaks appearing as doublets. The equalheight of the doublets suggests an equal molar ratio of theregioisomers. The peaks appearing at 152 ppm and 136 ppm are the carbonsα and β to the alkenyl nitro group, respectively.

FIG. 13 shows the spectrophotometric analysis of OA-NO₂. (A) Anabsorbance spectrum of OA-NO₂ from 200-450 nm was generated using 23 μMOA-NO₂ in phosphate buffer (100 mM, pH 7.4) containing 100 μM DTPA. Anabsorbance maximum at 270 nm was identified. (B) Extinction coefficientsfor OA-NO₂ and [¹³C]OA-NO₂ were determined by plotting absorbance (λ₂₇₀)vs. concentration, resulting in calculated values of ε=8.22 and 8.23cm⁻¹ mM⁻¹, respectively.

FIG. 14 shows the GC-MS analysis of synthetic OA-NO₂. (A) Methyl estersof the two synthetic OA-NO₂ regioisomers were generated as described inMethods and analyzed by EI GC MS/MS. The mixture was resolved using a 30m fused silica column and detected by total ion monitoring. The upperchromatogram shows partial resolution of the two regioisomers. Production analysis of each peak (B) revealed that the first and second elutingpeaks (37.41 and 37.59 min, respectively) each has a unique identifyingion: m/z 168 (peak 1) and m/z 156 (peak 2).

FIG. 15 shows the identification and characterization of synthetic andblood OA-NO₂ by HPLC ESI MS/MS. (A, left panels) OA-NO₂ and [¹³C]OA-NO₂were characterized by HPLC-ESI MS/MS. Nitrated oleic acid species wereseparated by HPLC and detected by acquiring MRM transitions consistentwith the loss of the alkenyl nitro group [M−HNO₂]⁻, m/z 326/279 and m/z344/297 for OA-NO₂ and [¹³C]OA-NO₂, respectively. (A, right panels)Concurrent to MRM detection, product ion analysis was performed togenerate identifying fragmentation patterns also used to characterize invivo OA-NO₂. The predominant product ions generated by collision-induceddissociation are identified in Table 3. (B) Total lipid extracts wereprepared from packed red cell and plasma fractions of venous blood anddirectly analyzed by mass spectrometry.

FIG. 16 shows the product ion analysis of fatty acidnitrohydroxy-adducts in urine. The presence of nitrohydroxy fatty acidsin urine was confirmed by product ion analysis run concomitant to MRMdetection. Structures of possible adducts are presented along with theirdiagnostic fragments and product ion spectra for (A) 18:1(OH)—NO₂, (B)18:2(OH)—NO₂ and (C) 18:3(OH)—NO₂. The 10-nitro regioisomer of18:1(OH)—NO₂ is present in urine, as evidenced by the intense peakcorresponding to m/z 171; also present are fragments consistent with the9-nitro regioisomer (m/z 202), loss of a nitro group (m/z 297) and water(m/z 326). 18:2(OH)—NO₂ also shows a predominant m/z 171 fragment, againconsistent with an oxidation product of LNO₂ nitrated at the 10-carbon(B). Diagnostic fragments for the three other potential regioisomerswere not apparent. Finally, multiple regioisomers of 18:3(OH)—NO₂ arepresent (C).

FIG. 17 shows that OA-NO₂ is a PPARγ agonist. (A) CV-1 cells transientlyco-transfected with a plasmid containing the luciferase gene under thecontrol of three tandem PPRE (PPRE×3 TK-Luciferase) and hPPARγ, hPPARαor hPPARδ expression plasmids showed all three PPARs were activated byOA-NO₂, with the relative activation of PPARγ>PPARδ>PPARα. All valuesare expressed as mean±SD (n=3). PPARγ activation was significantlydifferent from vehicle at 100 nM OA-NO₂, whereas PPARα and PPARδactivation were significantly different from vehicle at 300 nM and 1 μMOA-NO₂, respectively (P<0.05; Student's t test). (B) Nitrated oleic acidappears to be more potent than LNO₂ in the activation of PPARγ, with 1μM OA-NO₂ inducing similar activity as 3 μM LNO₂ versus control (P≦0.05;Student's t test). Activation of PPARγ was partially yet significantlyblocked using the PPARγ antagonist GW9662 (P≦0.05; Student's t test).(C) Equimolar concentrations of OA-NO₂ and LNO₂ (3 μM) were incubated in100 mM phosphate buffer. After 2 hr, only 25% of the initial OA-NO₂ haddegraded; ˜80% of LNO₂ degrades in the same time period, indicating agreat aqueous stability of OA-NO₂. All values are expressed as mean±SD(n>3).

FIG. 18 shows that OA-NO₂ induces adipogenesis in 3T3 L1 preadipocytes.PPARγ plays an essential role in the differentiation of adipocytes.3T3-L1 preadipocytes were treated with OA-NO₂, LNO₂, Rosiglitazone andcontrols (oleic acid, linoleic acid and DMSO) for two weeks. (A)Adipocyte differentiation was assessed both morphologically and via oilred O staining, which reveals the accumulation of intracellular lipids.Vehicle, oleic acid and linoleic acid did not induce adipogenesis, whileOA-NO₂ induced ˜60% of 3T3-L1 preadipocyte differentiation; LNO₂ induced˜30%, reflecting the greater potency of OA-NO₂. As the positive control,Rosiglitazone also induced PPARγ-dependent adipogenesis. (B) OA-NO₂ andRosiglitazone-induced preadipocyte differentiation resulted in theexpression of adipocyte-specific markers (PPARγ2 and aP2), an event notdetected for oleic acid.

FIG. 19 shows that OA-NO₂ induces [³H]-2-deoxy-D-glucose uptake indifferentiated 3T3L1 adipocytes. (A) PPARγ ligands induce glucose uptakein adipose tissue. To further define the functional significance ofOA-NO₂ as a PPARγ ligand, 3T3-L1 preadipocytes were differentiated toadipocytes and treated with OA-NO₂ or LNO₂ for two days prior toaddition of [³H]-2-deoxy-D-glucose. OA-NO₂ induced significant increasesin glucose uptake; these effects were paralleled by LNO₂ (P≦0.05;Student's t test). (B) The increases in glucose uptake induced bynitrated lipids and the positive control Rosiglitazone weresignificantly inhibited by the PPARγ-specific antagonist GW9662 (P≦0.05;Student's t test). All values are expressed as mean±SD (n=3).

FIG. 20 shows EPR and UV/visible spectroscopic detection of .NO releaseby LNO₂. (A) EPR spectral analysis of cPTIO (200 μM, red line) reductionto cPTI (black line) by LNO₂ (300 μM) decay during a 60 min decayperiod. (B) Differential spectra of oxymyoglobin (20 μM) oxidation byLNO₂ (200 μM). Spectra were repetitively recorded at 5 min intervals andshow the decrease in the 580 nm and 543 nm maxima (characteristic of theα and β visible band absorbance of the oxymyoglobin) and the increase in630 and 503 nm maxima characteristic of metmyoglobin. (C) .NO releaserate detected by oxymyoglobin (20 μM) oxidation in the presence ofdifferent concentrations of LNO₂. Values expressed as mean±SD of 2independent experiments repeated four times. (D) UV spectra of LNO₂taken every 10 min, revealing loss of the characteristic absorbance ofthe NO₂ group at 268 nm and the formation of a new chromophore at 320nm. (A, B and D) Spectra are representative of 3 independentexperiments.

FIG. 21 shows nitrite formation during LNO₂ decay. The time-dependentformation of NO₂ ⁻ during LNO₂ (initial concentration 200 μM)decomposition was measured in parallel with oxymyoglobin (20 μM)oxidation. Nitrite formation was measured in the absence ofoxymyoglobin. Values expressed as mean±SD of 3 independent experimentsrepeated three times.

FIG. 22 shows the pH dependency of .NO formation from LNO₂. The rate of.NO formation from 80 μM LNO₂ (detected using EPR spectroscopicmeasurement of cPTIO (80 μM) reduction) was determined in buffers withdifferent pHs.

FIG. 23 shows chemiluminescent detection of .NO release by LNO₂. A) LNO₂(5 and 10 mM) was incubated in a capped vial under aerobic conditionsfor 3 min and the gas phase injected into an O₃ chemiluminescencedetector. Additionally, known concentrations of DEA-NONOate (nM) (in 10mM NaOH) were added to a capped vial containing 0.5 M HCl and the gasphase was injected into the chemiluminescence detector. B) Phosphatebuffer (50 mM phosphate pH 7.4 containing 10 μM DTPA) was illuminatedwith a xenon arc lamp. .NO formation was examined by O₃-basedchemiluminescence after the injection of MeOH (20 μl), LNO₂ (4 nmol in20 μl MeOH, two additions made before and after sodium nitrite addition)and sodium nitrite (4 nmol in 20 μl phosphate buffer). C) Blood wasobtained by cardiac puncture of LPS-treated rats, red cells removed bycentrifugation and plasma samples treated as noted in ExperimentalProcedures. The following conditions were studied in panel C: (1) I₃ ⁻alone; (2) I₃ ⁻ plus sulfanilamide; (3) I₃ ⁻ plus sulfanilamide andHgCl₂, with 3.5 nmol LNO₂ treated with I₃ ⁻ plus sulfanilamide andHgCl₂, as for the corresponding plasma sample. Derived .NO was measuredby .NO chemiluminescence analysis. Traces are representative from threedifferent experiments.

FIG. 24 shows spectroscopic (EPR, UV and visible) analysis of micellarinhibition of LNO₂ decomposition and .NO release. In panels A-D, closeddiamonds represent conditions containing OTG and open diamonds representOG. A) .NO release from LNO₂ (80 μM) in the presence of different OTGand OG concentrations after 60 min, as measured by EPR detection ofcPTIO (80 μM) conversion to cPTI. The extent of cPTIO (80 μM) conversionto cPTI by known concentrations of proli-NONOate was utilized tocalculate yields of .NO. B) .NO release rate from LNO₂ (130 μM) in thepresence of different concentrations of OG and OTG, as measured byoxidation of oxymyoglobin (20 μM) to metmyoglobin. An extinctioncoefficient of 14.4 mM⁻¹ cm⁻¹ was used to calculate yields of .NO. C)Initial decomposition rates of LNO₂ (37 μM), measured at 268 nm, in thepresence of different concentrations of OTG and OG. D) Same as C, butrates of LNO₂ decomposition product formation at 320 nm were measured.Values are expressed as mean±SD of at least 3 independent experimentsrepeated three or four times.

FIG. 25 shows micellar and phosphatidylcholine-cholesterol liposomeinhibition of LNO₂ decomposition and .NO release. A) LNO₂ (200 μM)decomposition was measured in the absence and presence of 15 mg/ml OTGby mass spectrometry. B) Calculation of partition coefficient of LNO₂into OTG and OG micelles from data shown in FIG. 25D). C) LNO₂ (80μM)-dependent .NO formation was measured by cPTIO (200 μM) reduction tocPTI at different times in the presence of increasing liposomeconcentration (0-5 mg/ml).

FIG. 26 shows formation of L(OH)NO₂ from LNO₂. A) LNO₂ was incubated inthe presence (dotted line) or absence (solid line) of 15 mg/ml OTG,lipids were extracted and analyzed by ESI MS/MS. The presence of OTGinhibited LNO₂ decay as indicated by the MRM transition m/z 324/277 (A)and the formation of species with transitions m/z 342/171 and 342/295,which correspond to 9-hydroxy-10-nitro-12-octadecaenoic acidspecifically (B), and all L(OH)NO₂ regioisomers (C), respectively. Inthe absence of OTG, increased L(OH)NO₂ yields were formed. D) Structuresof possible nitrohydroxy adducts are presented along with theirdiagnostic fragments. E) Product ion spectra of L(OH)NO₂ showed twopredominant ions consistent with expected fragments shown in (D), m/z171 (9-hydroxy-10-nitro-12-octadecaenoic acid) and m/z 211(12-hydroxy-13-nitro-9-octadecaenoic acid

FIG. 27 shows mass spectrometric detection of LNO₂ decay products. LNO₂(500 μM) was incubated in aqueous buffer phosphate buffer (100 mMphosphate pH 7.4 containing 100 μM DTPA) for 0, 45 and 240 min (A-C,respectively). Decay products were CHCl₃-extracted and analyzed bydirect ESI MS/MS. Products were detected in the negative ion mode. The293 m/z ion corresponds to an expected Nef reaction product, aconjugated ketone; m/z 342 is consistent with the mass of vicinalnitrohydroxy linoleic acid; and m/z 340 and 356 represent the hydroxyand peroxy derivatives of LNO₂, respectively.

FIG. 28 shows Scheme 1, Hydrophobic regulation of LNO₂ decomposition and.NO release in lipid bilayers and micelles. The partitioning of LNO₂into different cell compartments is in part governed by its partitioncoefficient (K˜1500). LNO₂ may also be stabilized and placed in“reserve”, in terms of attenuating .NO-mediated cell signalingcapabilities, by esterification into complex lipids of membranes orlipoproteins. Alternatively, LNO₂ derivatives of complex lipids can beformed by direct nitration of esterified unsaturated fatty acids. Duringinflammatory conditions or in response to other stimuli, LNO₂ may bereleased from complex lipids by A₂-type phospholipases or esterases;thus mobilizing “free” LNO₂ that can in turn diffuse to exertreceptor-dependent signaling actions or undergo decay reactions torelease .NO.

FIG. 29 shows Scheme 2, possible mechanisms for NO formation by LNO₂.(Stage 1) Due to the strong electrophilic nature of the carbon adjacentto the nitroalkene and the acidity of its bound hydrogen, the vicinalnitrohydroxy fatty acid derivative is in equilibrium with thenitroalkene. (Stage 2) The mechanism of .NO release from LNO₂ can resultfrom the formation of a nitroso intermediate formed during aqueous LNO₂decay. This nitroso intermediate is expected to have an especially weakC—N bond, easily forming .NO and a radical stabilized by conjugationwith the alkene and stabilized by the OH group, a moiety known tostabilize adjacent radicals.

FIG. 30 shows that nitroalkenes potently activate p-JNK and p-c-Junprotein kinases by stimulating their phosphorylation. The activation ofthese cell signaling mediators will profoundly impact on cellinflammatory responses, proliferation and differentiation. This exampleshows human lung epithelial cell responses of p-JNK and p-c-Jun.

FIG. 31 shows that nitroalkenes activate the ERK MAPK pathway in humanlung epithelial cells, as shown by a dramatic increase in ERKphosphorylation (e.g., activation) and the phosphorylation of itsdownstream target signaling protein, pELK. The activation of these cellsignaling mediators will profoundly impact on cell inflammatoryresponses, proliferation and differentiation. This example shows humanlung epithelial cell responses of p-JNK and p-c-Jun.

FIG. 32 shows that nitrolinoleate (LNO₂), and not the control fatty acidlinoleate (LA), inhibits activity of NF-kB pathways as indicated by a)luciferase-linked NFkB-response element reporter assay in response tothe inflammatory mediator TNFa and b) direct analysis of the degradationf the NFkB inhibitor protein, IkB in response to the inflammatorymediator E. coli LPS.

FIG. 33 shows the mass spectra of nitro/hydroxy fatty acid.

FIG. 34 shows the mass spectrum of the Michael addition product betweennitro linoleate and glutathione.

DETAILED DESCRIPTION

Before the present compounds, compositions, and/or methods are disclosedand described, it is to be understood that the aspects described beloware not limited to specific compounds, synthetic methods, or uses assuch may, of course, vary. It is also to be understood that theterminology used herein is for the purpose of describing particularaspects only and is not intended to be limiting.

In this specification and in the claims that follow, reference will bemade to a number of terms that shall be defined to have the followingmeanings:

It must be noted that, as used in the specification and the appendedclaims, the singular forms “a,” “an” and “the” include plural referentsunless the context clearly dictates otherwise. Thus, for example,reference to “a pharmaceutical carrier” includes mixtures of two or moresuch carriers, and the like.

“Optional” or “optionally” means that the subsequently described eventor circumstance can or cannot occur, and that the description includesinstances where the event or circumstance occurs and instances where itdoes not. For example, the phrase “optionally substituted lower alkyl”means that the lower alkyl group can or can not be substituted and thatthe description includes both unsubstituted lower alkyl and lower alkylwhere there is substitution.

Ranges may be expressed herein as from “about” one particular value,and/or to “about” another particular value. When such a range isexpressed, another aspect includes from the one particular value and/orto the other particular value. Similarly, when values are expressed asapproximations, by use of the antecedent “about,” it will be understoodthat the particular value forms another aspect. It will be furtherunderstood that the endpoints of each of the ranges are significant bothin relation to the other endpoint, and independently of the otherendpoint.

References in the specification and concluding claims to parts byweight, of a particular element or component in a composition orarticle, denotes the weight relationship between the element orcomponent and any other elements or components in the composition orarticle for which a part by weight is expressed. Thus, in a compoundcontaining 2 parts by weight of component X and 5 parts by weightcomponent Y, X and Y are present at a weight ratio of 2:5, and arepresent in such ratio regardless of whether additional components arecontained in the compound.

A weight percent of a component, unless specifically stated to thecontrary, is based on the total weight of the formulation or compositionin which the component is included.

Variables such as R¹-R¹⁶ used throughout the application are the samevariables as previously defined unless stated to the contrary.

By “subject” is meant an individual. The subject can be a mammal such asa primate or a human. The term “subject” can include domesticatedanimals including, but not limited to, cats, dogs, etc., livestock(e.g., cattle, horses, pigs, sheep, goats, etc.), and laboratory animals(e.g., mouse, rabbit, rat, guinea pig, etc.).

By “contacting” is meant an instance of exposure by close physicalcontact of at least one substance to another substance. For example,contacting can include contacting a substance, such as a pharmacologicagent, with a cell. A cell can be contacted with a test compound, forexample, a nitrated lipid, by adding the agent to the culture medium (bycontinuous infusion, by bolus delivery, or by changing the medium to amedium that contains the agent) or by adding the agent to theextracellular fluid in vivo (by local delivery, systemic delivery,intravenous injection, bolus delivery, or continuous infusion). Theduration of contact with a cell or group of cells is determined by thetime the test compound is present at physiologically effective levels orat presumed physiologically effective levels in the medium orextracellular fluid bathing the cell.

“Treatment” or “treating” means to administer a composition to a subjector a system with an undesired condition (e.g., inflammation) or at riskfor the condition. The condition can include a disease or apredisposition to a disease. The effect of the administration of thecomposition to the subject can have the effect of but is not limited toreducing or preventing the symptoms of the condition, a reduction in theseverity of the condition, or the complete ablation of the condition.

By “effective amount” is meant a therapeutic amount needed to achievethe desired result or results, e.g., increasing the expression of agene, inhibiting Ca⁺² mobilization in a cell, inhibiting degranulationor CD11b expression in a neutrophil, etc.

Herein, “inhibition” or “suppression” means to reduce activity ascompared to a control. It is understood that inhibition or suppressioncan mean a slight reduction in activity to the complete ablation of allactivity. An “inhibitor” or “suppressor” can be anything that reducesthe targeted activity.

Herein, “induce” means initiating a desired response or result that wasnot present prior to the induction step. The term “potentiate” meanssustaining a desired response at the same level prior to thepotentiating step or increasing the desired response over a period oftime.

The term “alkyl group” as used herein is a branched or unbranchedsaturated hydrocarbon group of 1 to 24 carbon atoms, such as methyl,ethyl, n-propyl, isopropyl, n-butyl, isobutyl, t-butyl, pentyl, hexyl,heptyl, octyl, decyl, tetradecyl, hexadecyl, eicosyl, tetracosyl and thelike. A “lower alkyl” group is an alkyl group containing from one to sixcarbon atoms.

The term “alkenyl group” is defined as a branched or unbranchedhydrocarbon group of 2 to 24 carbon atoms and structural formulacontaining at least one carbon-carbon double bond.

The term “alkynyl group” is defined as a branched or unbranchedhydrocarbon group of 2 to 24 carbon atoms and a structural formulacontaining at least one carbon-carbon triple bond.

The term “ester” is represented by the formula —OC(O)R, where R can bean alkyl, alkenyl, or group described above.

R¹-R¹⁶ can, independently, possess two or more of the groups listedabove. For example, if R¹ is a straight chain alkyl group, one of thehydrogen atoms of the alkyl group can be substituted with an estergroup. Depending upon the groups that are selected, a first group may beincorporated within second group or, alternatively, the first group maybe pendant (i.e., attached) to the second group. For example, with thephrase “an alkyl group comprising an ester group,” the ester group maybe incorporated within the backbone of alkyl group. Alternatively, theester can be attached the backbone of the alkyl group. The nature of thegroup(s) that is (are) selected will determine if the first group isembedded or attached to the second group.

Disclosed are compounds, compositions, and components that can be usedfor, can be used in conjunction with, can be used in preparation for, orare products of the disclosed methods and compositions. These and othermaterials are disclosed herein, and it is understood that whencombinations, subsets, interactions, groups, etc. of these materials aredisclosed that while specific reference of each various individual andcollective combinations and permutation of these compounds may not beexplicitly disclosed, each is specifically contemplated and describedherein. For example, if a number of different nucleosides and polymericsubstrates are disclosed and discussed, each and every combination andpermutation of the nucleoside and the polymeric substrate arespecifically contemplated unless specifically indicated to the contrary.Thus, if a class of molecules A, B, and C are disclosed as well as aclass of molecules D, E, and F and an example of a combination molecule,A-D is disclosed, then even if each is not individually recited, each isindividually and collectively contemplated. Thus, in this example, eachof the combinations A-E, A-F, B-D, B-E, B-F, C-D, C-E, and C-F arespecifically contemplated and should be considered disclosed fromdisclosure of A, B, and C; D, E, and F; and the example combination A-D.Likewise, any subset or combination of these is also specificallycontemplated and disclosed. Thus, for example, the sub-group of A-E,B-F, and C-E are specifically contemplated and should be considereddisclosed from disclosure of A, B, and C; D, E, and F; and the examplecombination A-D. This concept applies to all aspects of this disclosureincluding, but not limited to, steps in methods of making and using thedisclosed compositions. Thus, if there are a variety of additional stepsthat can be performed it is understood that each of these additionalsteps can be performed with any specific embodiment or combination ofembodiments of the disclosed methods, and that each such combination isspecifically contemplated and should be considered disclosed.

I. Nitrated Lipids

In one aspect, the nitrated lipids described herein are lipidscomprising at least one nitro group (NO₂) covalently bonded to thelipid, wherein the nitrated lipid is substantially pure. The term“substantially pure” as defined herein is a nitrated lipid that existspredominantly as one species. In certain aspects, nitration of a lipidcan produce two or more nitration products. For example, the lipid canbe nitrated one or more times at different positions on the lipid. Theseare referred to as positional isomers. Additionally, if the lipidcontains a carbon-carbon double bond, the stereochemistry about thecarbon-carbon double bond can also vary. These are referred to asstereoisomers. The nitrated lipids described herein are substantiallyone compound (positional and stereoisomer). In one aspect, the nitratedlipid is 90%, 92%, 94%, 96%, 98%, 99%, 99.5%, or 100% one compound.

In one aspect, the nitrated lipids possess at least one allylic or vinylnitro group. The phrase “allylic nitro group” has the general formula—C═C—C—(NO₂). The phrase “vinyl nitro group” has the general formula—C═C—(NO₂). In one aspect, the nitrated lipid possesses only one allylicnitro group. In another aspect, the nitrated lipid possesses only onevinyl nitro group. In another aspect, the nitrated lipid possesses oneor more allylic nitro groups and/or one or more vinyl nitro groups.

Lipids known in the art can be nitrated using the techniques describedherein to produce nitrated lipids. In general, lipids useful forproducing the nitrated lipid include, but are not limited to, fats andfat derived materials. In one aspect, the nitrated lipid can include,but is not limited to, a nitrated fatty acid or ester thereof, anitrated fatty alcohol, or a nitrated sterol. In another aspect, thenitrated lipid can be a nitrated complex lipid. Examples of complexlipids include, but are not limited to, glycerolipids (e.g., compoundshaving a glycerol backbone including, but not limited to, phospholipids,glycolipids, monoglycerides, diglycerides, triglycerides) or cholesterol(e.g., cholesterols having fatty acids attached to it such ascholesterol linoleate). In one aspect, the nitrated lipid comprises afatty acid having at least one ester linkage [—O—C═O(R)], ether group(C—O—R) or vinyl ether group (C—O—C═C—R). Examples of lipids having atleast one ether group or vinyl ether group that can be nitrated aredepicted below in A and B, respectively.

wherein

-   -   R¹⁴ comprises C₁₆-C₂₂ alkyl, C₁₆-C₂₂ alkenyl, or C₁₆-C₂₂        alkynyl;    -   R¹⁵ comprises C₁-C₂₀ alkyl, C₁-C₂₀ alkenyl, or C₁-C₂₀ alkynyl;        and    -   R¹ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl.

In one aspect, the nitrated lipid is composed of a fatty acid having atleast one carbon-carbon double bond. In one aspect, the nitrated lipidcan be a nitrated fatty acid such as, for example, 14:1, 16:1, 18:1(oleic acid), 18:2 (linoleic acid), 18:3 (linolenic acid), 20:4(arachidonic acid), 22:6, or docosahexanoic acid, where the first numberindicates the carbon chain length of the fatty acid, and the secondnumber indicates the number of carbon-carbon double bonds present in thefatty acid.

In one aspect, the nitrated lipid can have the formula I

wherein

-   -   R¹ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl;    -   R², R³, R⁷, and R⁸ comprise, independently, hydrogen, NO₂, OH,        or OOH;    -   R⁴ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl;    -   wherein R⁴ comprises a terminal COOR⁶ group, wherein R⁶        comprises hydrogen, C₁-C₂₄ alkyl, or a pharmaceutically        acceptable counterion, wherein    -   R⁴ optionally comprises one or more NO₂, OH, or OOH groups;    -   R⁵ comprises hydrogen or R⁴ and R⁵ collectively forms        ═C(R⁹)(R¹⁰), wherein    -   R⁹ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl,        wherein R⁹ comprises a terminal COOR⁶ group, wherein R⁹        optionally comprises one or more NO₂, OH, or OOH groups;    -   R¹⁰ comprises hydrogen, NO₂, OH, or OOH; and    -   n is from 1 to 24;        wherein the nitrated lipid comprises at least one NO₂ group,        wherein the nitrated lipid is substantially pure. In this        aspect, the stereochemistry about the carbon-carbon double bond        is substantially cis (or Z) or substantially trans (or E).

In another aspect, the nitrated lipid can have the formula II

wherein R³ is trans or cis to R⁸, and R⁷ is trans or cis to R¹⁰,wherein the nitrated lipid is substantially pure. In this aspect, thestereochemistry about the carbon-carbon double bond is substantially cis(or Z) or substantially trans (or E). In one aspect, when the nitratedlipid has the formula II, R¹ comprises a C₄-C₁₀ alkyl group, R², R³, R⁸,and R¹⁰ are hydrogen, R⁷ is NO₂, and R⁹ comprises a C₆-C₁₂ alkyl group.This class of nitrated lipids is depicted below, where the nitratedlipid has one vinyl nitro group.

In a further aspect, when the nitrated lipid has the formula II, R³ andR⁸ are cis (Z) to one another, and R⁷ and R¹⁰ are cis (Z) to oneanother. In another aspect, the nitrated lipid is10-nitro-9-cis,12-cis-octadecadienoic acid, which is depicted in FIG.4C.

In another aspect, when the nitrated lipid has the formula II, R¹comprises a C₄-C₁₀ alkyl group, R², R³, R⁷, and R¹⁰ are hydrogen, R⁸ isNO₂, and R⁹ comprises a C₆-C₁₂ alkyl group. This class of nitratedlipids is depicted below, where the nitrated lipid has one vinyl nitrogroup.

In a further aspect, R³ and R⁸ are cis (Z) to one another and R⁷ and R¹⁰are cis (Z) to one another. In another aspect, the nitrated lipid is12-nitro-9-cis,12-cis-octadecadienoic acid, which is depicted in FIG.4C.

In another aspect, the nitrated lipid has the formula III

wherein

-   -   R¹ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl;    -   R² and R¹² comprise, independently, hydrogen, NO₂, OH, or OOH;        and    -   R¹³ comprises C₁-C₂₄ alkyl, C₁-C₂₄ alkenyl, or C₁-C₂₄ alkynyl,        wherein    -   wherein R¹³ comprises a terminal COOR⁶ group, wherein R⁶        comprises hydrogen, C₁-C₂₄ alkyl, or a pharmaceutically        acceptable counterion, wherein    -   R¹³ optionally comprises one or more NO₂, OH, or OOH groups;        wherein the compound comprises at least one NO₂ group,        wherein the nitrated lipid is substantially pure.

In another aspect, when the nitrated lipid has the formula III, R¹comprises a C₄-C₁₀ alkyl group, R² is hydrogen, R¹² is NO₂, and R¹³comprises a C₆-C₁₂ alkyl group. This class of nitrated lipids isdepicted below, where the nitrated lipid has one allylic nitro group.

In this aspect, R¹ comprises a C₄-C₁₀ alkyl group, R² is NO₂, R¹² ishydrogen, and R¹³ comprises a C₆-C₁₂ alkyl group. In one aspect, thenitrated lipid is 9-nitro-10,12-cis-octadecadienoic acid;9-nitro-10,12-trans-octadecadienoic acid;13-nitro-10,12-cis-octadecadienoic acid; or13-nitro-10,12-trans-octadecadienoic acid.

Methods for preparing the nitrated lipids described herein are describedbelow and in the Examples section.

II. Synthesis of Nitrated Lipids

Described herein are methods for preparing nitrated lipids. In oneaspect, the method comprises

(a) reacting an unsaturated lipid with a mercuric salt, a seleniumcompound, and a nitrating compound to produce a first intermediate, and

(b) reacting the first intermediate with an oxidant.

Any of the lipids described above can be used to produce the nitratedlipids described herein. In one aspect, any unsaturated lipid having atleast one carbon-carbon double bond can be used in this aspect toproduce nitrated lipids.

In one aspect, step (a) can be performed in situ without isolation ofthe first intermediate. In another embodiment, the first intermediatecan be trapped prior to step (b). The mercuric salt, a seleniumcompound, and a nitrating compound can be added in any order to theunsaturated lipid.

The selenium compound is any compound that is capable of reacting orinteracting with the unsaturated group present in the lipid. In oneaspect, when the unsaturated lipid possesses a carbon-carbon doublebond, the selenium compound can form a three-membered ring intermediate,which is depicted, for example, in FIG. 1. In one aspect, the seleniumcompound possesses a leaving group capable of reacting with the threering intermediate. Examples of leaving groups include, but are notlimited to, halides (e.g., F, Cl, Br, I) and esters (e.g., acetate). Notwishing to be bound by theory, it is believed that the leaving groupreacts with the three-membered ring intermediate to open the ring toproduce a selenium-leaving group intermediate. One example of thisintermediate is depicted in FIG. 1, where Br is the leaving group.Examples of selenium compounds useful herein include, but are notlimited to, PhSeBr, PhSeCl, PhSeO₂CCF₃, PhSeO₂H, or PhSeCN.

The mercuric salt used in the methods described herein can be anymercuric salt known in the art. Not wishing to be bound by theory, it isbelieved that the mercuric salt facilitates the formation of theselenium three-membered ring intermediate. In one aspect, the mercuricsalt comprises HgCl₂, Hg(NO₃)₂, or Hg(OAc)₂.

The nitrating compound is any compound that provides a source of NO₂ ⁻ions in solution. Not wishing to be bound by theory, it is believed thatNO₂ ⁻ reacts with the selenium intermediate formed upon the reactionbetween the unsaturated lipid and selenium compound by displacing theleaving group present in the intermediate. FIG. 1 depicts one aspect ofthis, where NO₂ ⁻ displaces Br⁻ by nucleophilic attack to produce aselenium/nitro intermediate. In one aspect, the nitrating compound canbe any nitrite salt. In another aspect, the nitrating compound comprisesNaNO₂ or AgNO₂.

The relative amounts of unsaturated lipid, selenium compound, mercuricsalt, and nitrating compound can vary depending upon the specificreagents that are selected and reaction conditions. In one aspect, anequimolar amount or slight excess thereof of selenium compound, mercuricsalt, and nitrating compound relative to the unsaturated lipid can beused. Step (a) is generally performed in a solvent, such as, forexample, a polar or unpolar organic solvent. Examples of solvents usefulherein include, but are not limited to, nitriles, ethers, esters,alkanes, alcohols, or combinations thereof. In one aspect, the solventused in step (a) can be THF/acetonitrile. Step (a) can be performed atvarious temperatures depending upon the starting materials that areselected. In one aspect, the step (a) can be performed at roomtemperature.

By varying the reaction conditions in step (a), it is possible toincrease the overall yield of the nitrated lipids as well as reduce thenumber of different types of nitrated lipids. In one aspect, the step(a) can be performed under anaerobic conditions. In another aspect, step(a) can be performed under anhydrous conditions. In a further aspect,step (a) is performed under anaerobic and anhydrous conditions.

After step (a), the intermediate that is produced is reacted with anoxidant to convert the intermediate to the nitrated lipid (step (b)).Not wishing to be bound by theory, the oxidant oxidizes the seleniumcompound to produce a selenium-oxo group, which rearranges to producethe nitrated lipid. One aspect of this mechanism is depicted in FIG. 1,whereupon oxidation, the selenium-oxo group rearranges to produce PhSeOHand a nitrated lipid possessing a vinyl nitro group. In one aspect, theoxidant comprises H₂O₂ or an organic hydroperoxide. The amount ofoxidant will vary depending upon the selection of the oxidant andreaction conditions. In one aspect, a 2-fold, 3-fold, 5-fold, 7-fold,9-fold, 10-fold, 15-fold, or 20-fold molar amount of oxidant is usedcompared to the molar amount of unsaturated lipid. In one aspect, step(b) is performed in an organic solvent at reduced temperature.

In one aspect, the unsaturated lipid comprises oleic acid, linoleicacid, linolenic acid, arachidonic acid, eicosapentaenoic acid, ordocosahexanoic acid, the mercuric salt comprises HgCl₂, the seleniumcompound comprises PhSeBr, the nitrating compound comprises NaNO₂, andthe oxidant comprises H₂O₂. In a further aspect, step (a) is performedunder anaerobic and anhydrous conditions.

After step (b), one nitrated lipid or mixture of two or more nitratedlipid positional or stereoisomers may be present depending upon reagentsand starting materials that are selected. In one aspect, if two or morenitrated lipids are present, each of the nitrated lipids can beseparated to produce substantially pure nitrated lipid. In one aspect,the mixture of two or more nitrated lipids can be separated bychromatography. For example liquid chromatography, thin layerchromatography or column chromatography using silicic acid, silical gelor other adsorbents useful for lipid separations, can be used toseparate the nitrated lipids. The solvent system used in this aspectwill vary depending upon the type and number of nitrated lipids to beseparated and can be determined by one of ordinary skill in the art.

Also described herein are methods for stabilizing nitrated lipids,comprising placing the nitrated lipid in a hydrophobic medium. Thenitrated lipids are generally stable oils and can be stored indefinitelyin organic solvents under anaerobic and anhydrous conditions and reducedtemperature. Alternatively, the salts of the nitrated lipids are stableand can be further processed into a formulation. Alternatively, the morestable nitrohydroxy derivative can be formed under alkaline conditions,which at more neutral pH reversibly yields the parent nitrated fattyacid. Alternatively, the nitrated lipids can be placed in detergentemulsions or liposomes, which can be later administered to a subject.

III. Nitro/Hydroxy Lipids and Synthesis Thereof

In one aspect, described herein are lipids comprising at least one nitrogroup and at least one hydroxyl group, wherein the compound issubstantially pure. In one aspect, the nitro group and the hydroxylgroup are on adjacent carbon atoms. In this aspect, the lipid containsthe fragment HO—C—C—NO₂. In another aspect, the nitro/hydroxy lipid hasthe formula X or XI

wherein R¹ comprises a C₁-C₂₄ alkyl group, a C₁-C₂₄ alkenyl group, orC₁-C₂₄ alkynyl group, and R⁹ comprises a terminal COOR⁶ group, whereinR⁶ comprises hydrogen, C₁-C₂₄ alkyl, or a pharmaceutically acceptablecounterion, and p is from 1 to 12. In a further aspect, thenitro/hydroxy lipid has the formula XII or XIII

wherein R¹ comprises a C₁-C₂₄ alkyl group, a C₁-C₂₄ alkenyl group, orC₁-C₂₄ alkynyl group, and R⁹ comprises a terminal COOR⁶ group, whereinR⁶ comprises hydrogen, C₁-C₂₄ alkyl, or a pharmaceutically acceptablecounterion, and m is from 0 to 12.

Any of the nitrated lipids can be converted to the correspondingnitro/hydroxyl compound. In one aspect, the nitrated lipid is placed inan aqueous base. Not wishing to be bound by theory, it is believed thatthe nitrated lipid undergoes a Michael addition reaction with waterfollowed by deprotonation to produce the nitro/hydroxyl lipid. Thismechanism is depicted below.

The nitro/hydroxyl lipid can be subsequently isolated by solventextraction followed by purification using techniques known in the art(e.g., HPLC or thin layer chromatography).IV. Methods of Use

Delivery

As used throughout, administration of any of the nitrated lipidsdescribed herein can occur in conjunction with other therapeutic agents.Thus, the nitrated lipids can be administered alone or in combinationwith one or more therapeutic agents. For example, a subject can betreated with a nitrated lipid alone, or in combination withchemotherapeutic agents, antibodies, antivirals, steroidal andnon-steroidal anti-inflammatories, conventional immunotherapeuticagents, cytokines, chemokines, and/or growth factors. Combinations maybe administered either concomitantly (e.g., as an admixture), separatelybut simultaneously (e.g., via separate intravenous lines into the samesubject), or sequentially (e.g., one of the compounds or agents is givenfirst followed by the second). Thus, the term “combination” or“combined” is used to refer to either concomitant, simultaneous, orsequential administration of two or more agents. Furthermore, two ormore nitrated lipids described herein can be administered to a subjectconcomitantly, simultaneously, or sequentially.

The nitrated lipids can be administered in a number of ways depending onwhether local or systemic treatment is desired, and on the area to betreated. Administration may be topically (including opthamalically,vaginally, rectally, intranasally), orally, by inhalation, orparenterally, for example by intravenous drip, subcutaneous,intraperitoneal or intramuscular injection. The disclosed compounds canbe administered intravenously, intraperitoneally, intramuscularly,subcutaneously, intracavity, transdermally, intratracheally,extracorporeally, or topically (e.g., topical intranasal administrationor administration by inhalant). As used herein, “topical intranasaladministration” means delivery of the compositions into the nose andnasal passages through one or both of the nares and can comprisedelivery by a spraying mechanism or droplet mechanism, or throughaerosolization of the nucleic acid or vector. The latter can beeffective when a large number of subjects are to be treatedsimultaneously. Administration of the compositions by inhalant can bethrough the nose or mouth via delivery by a spraying or dropletmechanism. Delivery can also be directly to any area of the respiratorysystem (e.g., lungs) via intubation.

Parenteral administration of the composition, if used, is generallycharacterized by injection. Injectables can be prepared in conventionalforms, either as liquid solutions or suspensions, solid forms suitablefor solution of suspension in liquid prior to injection, or asemulsions. A more recently revised approach for parenteraladministration involves use of a slow release or sustained releasesystem such that a constant dosage is maintained. See, e.g., U.S. Pat.No. 3,610,795, which is incorporated by reference herein in its entiretyfor the methods taught.

The compositions may be in solution or in suspension (for example,incorporated into microparticles, liposomes, or cells). Thesecompositions may be targeted to a particular cell type via antibodies,receptors, or receptor ligands. The following references are examples ofthe use of this technology to target specific proteins to given tissue(Senter, et al., Bioconjugate Chem., 2:447-451, (1991); Bagshawe, K. D.,Br. J. Cancer, 60:275-281, (1989); Bagshawe, et al., Br. J. Cancer,58:700-703, (1988); Senter, et al., Bioconjugate Chem., 4:3-9, (1993);Battelli, et al., Cancer Immunol. Immunother., 35:421-425, (1992);Pietersz and McKenzie, Immunolog. Reviews, 129:57-80, (1992); andRoffler, et al., Biochem. Pharmacol, 42:2062-2065, (1991)). Vehiclessuch as “stealth” and other antibody conjugated liposomes (includinglipid mediated drug targeting to colonic carcinoma), receptor mediatedtargeting of DNA through cell specific ligands, lymphocyte directedtumor targeting, and highly specific therapeutic retroviral targeting ofmurine glioma cells in vivo. In general, receptors are involved inpathways of endocytosis, either constitutive or ligand induced. Thesereceptors cluster in clathrin-coated pits, enter the cell viaclathrin-coated vesicles, pass through an acidified endosome in whichthe receptors are sorted, and then either recycle to the cell surface,become stored intracellularly, or are degraded in lysosomes. Theinternalization pathways serve a variety of functions, such as nutrientuptake, removal of activated proteins, clearance of macromolecules,opportunistic entry of viruses and toxins, dissociation and degradationof ligand, and receptor-level regulation. Many receptors follow morethan one intracellular pathway, depending on the cell type, receptorconcentration, type of ligand, ligand valency, and ligand concentration.Molecular and cellular mechanisms of receptor-mediated endocytosis hasbeen reviewed (Brown and Greene, DNA and Cell Biology 10:6, 399-409(1991)).

The exact amount of the compositions required will vary from subject tosubject, depending on the species, age, weight and general condition ofthe subject, the severity of the disorder being treated, the particularnucleic acid to be targeted, its mode of administration and the like.Thus, it is not possible to specify an exact amount for everycomposition. However, an appropriate amount can be determined by one ofordinary skill in the art using only routine experimentation given theteachings herein. In one aspect, the amount of nitrated lipid that isadministered can be from 1 nM to 1 mM, 10 nM to 1 mM, 20 nM to 1 mM, 50nM to 1 mM, 100 nM to 1 mM, 200 nM to 1 mM, 300 nM to 1 mM, or 500 nM to1 mM. The time at which the nitrated lipids can be administered willalso vary depending upon the subject, the disorder, mode ofadministration, etc. The nitrated lipid can be administered to thesubject prior to the onset of inflammation or during a time when thesubject is experiencing inflammation. In one aspect, the nitrated lipidcan be administered within 24 hours, 20 hours, 16 hours, 12 hours, 8hours, 4 hours, 2 hours, 1 hour, or 30 minutes before inflammationoccurs or 10 hours, 20 hours, 30 hours, 40 hours, 60 hours, 80 hours,100 hours, or 120 hours after the onset of the inflammation.

Pharmaceutically Acceptable Carriers

The nitrated lipids can be used therapeutically in combination with apharmaceutically acceptable carrier. Pharmaceutical carriers are knownto those skilled in the art. These most typically would be standardcarriers for administration of drugs to humans, including solutions suchas sterile water, saline, and buffered solutions at physiological pH.The compositions can be administered intramuscularly or subcutaneously.Other compounds will be administered according to standard proceduresused by those skilled in the art.

Pharmaceutical compositions may include carriers, thickeners, diluents,solvents, buffers, preservatives, surface active agents and the like inaddition to the molecule of choice. Pharmaceutical compositions may alsoinclude one or more active ingredients such as antimicrobial agents,anti-inflammatory agents, anesthetics, and the like.

Preparations for parenteral administration include sterile aqueous ornon-aqueous solutions, suspensions, and emulsions. Examples ofnon-aqueous solvents are propylene glycol, polyethylene glycol,vegetable oils such as olive oil, and injectable organic esters such asethyl oleate. Aqueous carriers include water, alcoholic/aqueoussolutions, emulsions or suspensions, including saline and bufferedmedia. Parenteral vehicles include sodium chloride solution, Ringer'sdextrose, dextrose and sodium chloride, lactated Ringer's, or fixedoils. Intravenous vehicles include fluid and nutrient replenishers,electrolyte replenishers (such as those based on Ringer's dextrose), andthe like. Preservatives and other additives may also be present such as,for example, antimicrobials, anti-oxidants, chelating agents, and inertgases and the like.

Formulations for topical administration may include ointments, lotions,creams, gels, drops, suppositories, sprays, liquids and powders.Conventional pharmaceutical carriers, aqueous, powder or oily bases,thickeners and the like may be necessary or desirable.

Compositions for oral administration include powders or granules,suspensions or solutions in water or non-aqueous media, capsules,sachets, or tablets. Thickeners, flavorings, diluents, emulsifiers,dispersing aids or binders may be desirable.

Some of the nitrated lipids may potentially be administered as apharmaceutically acceptable acid- or base-addition salt, formed byreaction with inorganic acids such as hydrochloric acid, hydrobromicacid, perchloric acid, nitric acid, thiocyanic acid, sulfuric acid, andphosphoric acid, and organic acids such as formic acid, acetic acid,propionic acid, glycolic acid, lactic acid, pyruvic acid, oxalic acid,malonic acid, succinic acid, maleic acid, and fumaric acid, or byreaction with an inorganic base such as sodium hydroxide, ammoniumhydroxide, potassium hydroxide, and organic bases such as mono-, di-,trialkyl and aryl amines and substituted ethanolamines. In anotheraspect, the nitrated lipid is in the form of the sodium or potassiumsalt. In another embodiment, the nitrated lipids can be converted to thecorresponding pharmaceutically-acceptable ester such as, for example,the methyl ester.

Therapeutic Uses

The methods described herein contemplate the use of single or mixturesof two or more nitrated or nitro/hydroxyl lipids. In one aspect,disclosed are methods for reducing or preventing inflammation in asubject with inflammation or at risk for inflammation, comprisingadministering an effective amount of any of the nitrated lipidsdescribed herein, wherein the nitrated lipid reduces or prevents theinflammation in the subject. Examples of inflammation include, but arenot limited to, pulmonary inflammation, vascular inflammation, renalinflammation, inflammation of the central nervous system, hepaticinflammation, or splanchnic inflammation. The inflammation can beassociated with an inflammatory disease including, but not limited to,systemic lupus erythematosus, Hashimoto's disease, rheumatoid arthritis,graft-versus-host disease, Sjögren's syndrome, pernicious anemia,Addison disease, scleroderma, Goodpasture's syndrome, Crohn's disease,autoimmune hemolytic anemia, myasthenia gravis, multiple sclerosis,Alzheimer's disease, amyotrophic lateral sclerosis, Basedow's disease,thrombopenia purpura, insulin-dependent diabetes mellitus, allergy;asthma, inflammatory bowel disease, cancer, ulcerative colitis,scleroderma, cardiomyopathy, atherosclerosis, hypertension, sickle celldisease, or respiratory distress syndrome of neonate and adults. Inanother aspect, the inflammation can be caused by an organtransplantation, respiratory distress, ventilator induced lung injury,ischemia reperfusion, hemorrhagic shock, or sepsis. In one aspect, whenthe pulmonary inflammation is caused by respiratory distress or sepsis,the nitrated lipids can reduce or prevent the accumulation of alveolarfluid in a subject.

In one aspect, the nitrated lipids described herein can modulate theexpression or activity of one or more nucleic acids that encode one ormore inflammatory-related or cell signaling polypeptides. The term“modulate” is defined herein as the ability of the nitrated lipid todecrease or increase the expression or activity relative to a control.The “control” can be either the amount of expression or activity in theabsence of a nitrated lipid, or in the presence of solvent ornon-nitrated parent lipid used for the synthesis of the nitrated lipidderivative. Alternatively, the “control” can be the amount of expressionor activity before or after the period use (i.e., administration of thenitrated lipid). The term “inflammatory-related polypeptide” is definedherein as any polypeptide that can induce or potentiate an inflammatoryresponse. Examples of genes that can be modulated by the nitrated lipidsdescribed herein and inflammatory-related polypeptides include, but arenot limited to, prostaglandin H synthase-2, gamma glutamyl cysteinesynthase, low density lipoprotein receptor, vascular endothelial growthfactor, tocopherol binding protein, a heat shock protein (e.g., 10, 40,60, 70, 90 kD), prostaglandin receptor EP4, a protein tyrosinephosphatase, a Ca, Na, or K ATPase, a G-protein signaling regulator(e.g., 24 kD), a vasoactive intestinal peptide, a guanine nucleotideexchange factor, a bone morphogenetic protein, an aromatic-induciblecytochrome P450s, an ADP-ribosyltransferase, endothelin-1, a vascularcell adhesion molecule-1, an intercellular adhesion molecule-1, a tumornecrosis factor alpha receptor, an interleukin 1 beta receptor, atransforming growth factor beta receptor, an advanced glycationendproduct-specific receptor, a cAMP phosphodiesterase, a cGMPphosphodiesterase, a cyclin-dependent kinase, cathepsins B and D, aconnective tissue growth factor, actin, myosin, a tubulin gene, a c-srctyrosine kinase, an insulin growth factor binding protein, acysteine-rich angiogenic inducer 61, thrombospondin-1, acadherin-associated protein beta, heme oxygenase-1, one or more of thegenes listed in Table 5, one more of the genes listed in Table 6, one ormore of the genes listed in Table 7, or a combination thereof. In oneaspect, the nitrated lipid can modulate the nucleic acid 1.5 fold, 2fold, 3-fold, 5-fold, 10-fold, 20-fold, 30-fold, 40-fold, 50-fold,75-fold, 100-fold, 125-fold, 150-fold, or greater.

The nitrated lipids described herein can be used to mediate receptors ina cell in order to reduce or prevent inflammation in a subject. In oneaspect, described herein are methods for inducing or potentiatingperoxisome proliferator activated receptor (PPAR) activity, comprisingcontacting a cell comprising at least one PPAR receptor with one or morenitrated lipids under conditions that allow the compound to induce orpotentiate the activity of the PPAR receptor. The PPAR receptor can beα, δ, or γ. Alternatively, other lipid receptors that nitrated lipidscan bind to, be transported by, and mediate include the family of fattyacid binding proteins, G protein-coupled receptors and cis-retinoic acidbinding protein. Not wishing to be bound by theory, it is believed thatactivation of cell receptors such as, for example, PPAR, can modulatethe tissue expression and activity of inflammatory-related genes. Anumber of cell-types can be contacted with the nitrated lipids describedherein in order to reduce or prevent inflammation in a subject. Examplesof such cells include, but are not limited to the constituent cells oflung, airways, nasal passages, eyes, auditory system, liver, spleen,kidney, intestine, colon, genito-urinary tract, heart, brain, spinalcord, muscle, bone, connective tissue, blood and reticuloendothelialsystem and nervous tissue.

In another aspect, the nitrated lipids described herein can induce theexpression or potentiate the activity of an inflammatory-relatedpolypeptide when a cell comprising at least one nucleic acid thatencodes the inflammatory-related polypeptide is contacted with anitrated lipid under conditions that allow the compound to induce theexpression or potentiate the activity of the inflammatory-relatedpolypeptide. In one aspect, the following expressions or activities canbe induced or potentiated with the nitrated lipids described herein:

-   1. Modification of downstream signaling regulated by small    G-proteins.-   2. Increased phosphorylation and activation of c-Jun N-terminal    kinase.-   3. Increased the phosphorylation and activation of the transcription    factor c-Jun.-   4. Modification of activator protein-1 binding and activator    protein-1 mediated gene expression-   5. Increased phosphorylation and activation of extracellular-signal    regulated kinase.-   6. Increased phosphorylation and activation of the transcription    factor Elk-1.-   7. Modification of binding to serum response element (SRE) and SRE    mediated gene expression.-   8. Increased synthesis of the transcription factor c-Fos.-   9. Affect the nuclear translocation of the transcription factor    Nrf-2 (Nuclear factor erythroid 2 related factor 2).-   10. Modification of the electrophilic response element (also known    as antioxidant response element) binding and the electrophilic    response element mediated gene expression.-   11. Modification of p38 mitogen activated protein kinase activation.-   12. Modification of the nuclear translocation of the transcription    factor p65.-   13. Modification of nuclear factor-kappa B binding and nuclear    factor kappa B mediated gene expression.

In other aspects, any of the nitrated lipids described herein can reducea cell's response to an inflammatory stimulus. In one aspect, the cellcan be a neutrophil, monocyte, or macrophage. For example, the nitratedlipids can inhibit neutrophil, monocyte, or macrophage degranulation(e.g., azurophilic) or release of hydrolases and proteases followingdegranulation, neutrophil, monocyte, or macrophage O₂.⁻ formation,expression of CD11b expression in a neutrophil, monocyte, or macrophage,and fMLP-induced Ca⁺² influx in a neutrophil, monocyte, or macrophageupon contact of the neutrophil, monocyte, or macrophage with thenitrated lipid. In these aspects, the neutrophil, monocyte, ormacrophage can be contacted with the nitrated lipid in vivo, in vitro,or ex vivo.

In one aspect, described herein are methods for regulating the activityof a protein kinase signaling pathway, comprising reacting a kinase witha nitro lipid described herein. In another aspect, the nitrated lipidsdescribed herein can regulate the activity of a thiol-dependent enzymein a cell by contacting the cell with a nitrated lipid of the presentinvention. The cell can be contacted with the nitrated lipid, in vivo,in vitro, or ex vivo. For example, nitroalkenes inhibit enzymes thatdepend on thiols as catalytic residues. Not wishing to be bound bytheory, it is believed that the thiol can react with the nitrated lipidvia a Michael addition reaction. Via this property, nitroalkenes alsoserve a potent stimuli of thiol modification-dependent protein kinasesand their downstream cell signaling pathways. In this regard, a varietyof cell protein kinases are activated by nitroalkenes and the associatedthiol-dependent phosphoprotein phosphatases can also be inhibited bythiol alkylation. In one aspect, any thiol-dependent structural, cellsignaling and catalytic protein can be regulated by the nitrated fattyacid compounds described herein.

In another aspect, the nitrated lipids described herein can controlprotein trafficking in cells by serving as a thiol-attached hydrophobicmembrane trafficking agent for proteins to which they attach. Thus, thealkylation of proteins by membrane-avid nitrated fatty acids willfacilitate the localization of proteins containing hydrophobic nitratedfatty acid-amino acid adducts to cytosol, plasma membrane and organelle(nucleus, endoplasmaic reticulum, mitochondrial, golgi, secretoryvesicles) membranes.

In another aspect, the nitrated lipids described herein can inhibitplatelet function in a subject upon administration of the nitrated lipidto the subject. In one aspect, the nitrated lipids can inhibit thrombin-or other stimuli-induced platelet aggregation by attenuatingcAMP-dependent Ca⁺² mobilization and activation of the phosphorylationof vasodilator-stimulated phosphoprotein (VASP).

In another aspect, the nitrated lipids described herein can be used toinduce or potentiate tissue repair in a subject suffering frominflammation. Not wishing to be bound by theory, it is believed that thenitrated lipids can down regulate events that result in inflammation orthe impairment of vascular function and blood flow.

In another aspect, the nitrated lipids described herein can promotesatiety in a subject upon administration of the nitrated lipid to thesubject.

In another aspect, the nitrated lipids described herein can treat cancerupon administering an effective amount of the nitrated lipid to thesubject. Not wishing to be bound by theory, it is believed that thenitrated lipids directly stimulate tumor cell killing and potentiate thekilling of tumor cells by standard chemotherapeutic drugs. The cellnecrosis and apoptosis-inducing activity is believed to be the result ofnitrated lipid/PPAR ligand activity (e.g., PPAR activation), as well asby stimulating other cell signaling pathways noted above that mediatecell growth, cell differentiation and death signaling pathways.

In another aspect, the nitrated lipids set forth herein can act asnitric oxide (NO) donors, as described in the Examples. Therefore, thenitrated lipids described herein can be used to administer NO to asubject and/or treat a NO-related condition in a subject. Theseconditions include, but are not limited to, atherosclerosis, myocardialinfarction, peripheral vascular disease, coronary artery diseases, heartfailure, stroke, essential hypertension, diabetes mellitus,pre-eclampsia, erectile dysfunction, impotence, diabetic nephropathy,inflammatory glomerular diseases, acute renal failure, chronic renalfailure, inflammation, bacterial infection, septic shock, respiratorydistress syndromes, arthritis, cancer, impetigo, epidermolysis bullosa,eczema, neurodermatitis, psoriasis, pruritis, erythema, hidradenitissuppurativa warts, diaper rash and jock itch.

In another aspect, described herein are methods for detectinginflammation in a subject, comprising (a) measuring the amount of anitrated lipid present in the subject and (b) comparing the amount ofnitrated lipid in the subject to the amount of nitrated lipid present ina subject that is not experiencing any inflammation. In one aspect,patients with cardiovascular disease have increased amounts of nitratedfatty acid. Based on the presence of nitrated lipids, the detection ofincreased levels of nitrated fatty acids in blood, tissues and bodilyfluids can serve to diagnose the occurrence, progression and/orresolution of the inflammatory process.

EXAMPLES

The following examples are put forth so as to provide those of ordinaryskill in the art with a complete disclosure and description of how thecompounds, compositions, and methods described and claimed herein aremade and evaluated, and are intended to be purely exemplary and are notintended to limit the scope of what the inventors regard as theirinvention. Efforts have been made to ensure accuracy with respect tonumbers (e.g., amounts, temperature, etc.) but some errors anddeviations should be accounted for. Unless indicated otherwise, partsare parts by weight, temperature is in ° C. or is at ambienttemperature, and pressure is at or near atmospheric. There are numerousvariations and combinations of reaction conditions, e.g., componentconcentrations, desired solvents, solvent mixtures, temperatures,pressures and other reaction ranges and conditions that can be used tooptimize the product purity and yield obtained from the describedprocess. Only reasonable and routine experimentation will be required tooptimize such process conditions.

Example 1 Materials

Linoleic acid was purchased from Nu-Check Prep (Elysian, Minn.).Phenylselenium bromide, HgCl₂, NaNO₂, anhydrous tetrahydrofuran,N,N-diisopropylethylamine (99.5%) and acetonitrile were obtained fromSigma/Aldrich (St Louis, Mo.). Silica gel HF thin layer chromatography(TLC) plates (250 μm) were from Analtech (Newark, Del.).Pentafluorobenzyl bromide and methanolic BF₃ was from Pierce (Rockford,Ill.). Solvents used in synthesis were HPLC grade or better and werepurchased from Fisher Scientific (Fairlawn, N.J.). Solvents used formass spectrometric analyses from Burdick and Jackson (Muskigon, Mich.).[¹³C]Linoleic acid was from Spectra Stable Isotopes (Columbia, Md.) and[¹⁵N]NaNO₂ was from Cambridge Isotope Laboratories, Inc. (Andover,Mass.). [¹⁴N]LNO₂, [¹³C]LNO₂ and [¹⁵N]LNO₂ positional isomers weresynthesized as described previously for nitrated fatty acids.

LNO₂ Synthesis.

Linoleic acid/HgCl₂/phenylselenium bromide/NaNO₂ (1:1.3:1:1, mol/mol)were combined in THF/acetonitrile (1:1, v/v) with a final concentrationof 0.15 M linoleic acid. Care was taken to use anhydrous solvents, dryglassware and reagents that had been dried in vacuo over phosphoruspentoxide. The reaction mixture was stirred (4 h, 25° C.) followed bycentrifugation to sediment the precipitate. The supernatant wasrecovered, the solvent evaporated in vacuo, the product mixtureredissolved in THF (original volume) and the temperature reduced to 0°C. A 10-fold molar excess of H₂O₂ was slowly added with stirring to themixture then allowed to rest in an ice bath for 20 min followed by agradual warming to room temp (45 min). The product mixture was extractedwith equal parts saturated NaCl and diethyl ether, the organic phasecollected, the solvent removed in vacuo and the lipid products wereresolvated in CH₂Cl₂/CH₃OH (4:1, v/v). A mixture of LNO₂ positionalisomers were initially separated from the product mixture by preparativeTLC using silica gel HF plates developed twice in a solvent systemconsisting of hexane/ether/acetic acid (70:30:1, v/v). Regions of silicacontaining LNO₂ were scraped, extracted (29) and stored in CH₃OH underargon at −80° C. Under these conditions, purified nitrated linoleic acidis stable for >3 months. Large scale purification of the individualpositional isomers was performed by preparative HPLC using a 250×21.2 mmC18 Phenomenex Luna column (5 μm particle size). Lipids were eluted fromthe column using a gradient solvent system consisting of A (H₂Ocontaining 0.1% NH₄OH) and B (CNCH₃ containing 0.1% H₂O) under thefollowing conditions: 20-80% B (linear increase, 45 min), 80% B (2 min),20% B (5 min). Fractions were collected as positional isomers eluted,the solvent removed in vacuo and the lipids were stored in CH₃OH underargon at −80° C. Stable isotopes of LNO₂, specifically [¹³C]LNO₂ and[¹⁵N]LNO₂, were synthesized as above, except that [¹³C]linoleic acid or[¹⁵N]NaNO₂ were substituted in the synthetic scheme. FIG. 1 depicts asynthetic scheme for producing nitrated lipids. The process describedabove significantly increased the purity and yield of fatty acid allylicnitration products, facilitating structural resolution of specific LNO₂positional isomers and future cell signaling studies (22, 32, 33).Preparative TLC permitted the initial resolution of nitrated fatty acidsfrom starting materials and oxidized linoleic acid species (FIG. 2). Theprincipal C-10 and C-12 positional isomers have R_(f) values of 0.45 and0.50, respectively. The changes in synthetic approaches, use ofanhydrous reagents and the execution of nitrosenylation reaction stepsunder a nitrogen atmosphere resulted in an increase in yield of nitratedlinoleic acid products from 4% to 56%.

In an alternate aspect, nitrated fatty acids can be separated andpurified from oils such as, for example, fish oil, soy bean oil, andolive oil. For example, silica gel and silicic acid columns can be usedto fractionate the nitro-fatty acids, followed by preparative TLC orHPLC to separate the different nitrated fatty acids. Nitro-oleate is thepredominant (>90%) marine and plant-nitroalkene.

Spectral Analysis of LNO₂.

Initial concentrations of synthetic LNO₂ preparations were measured bychemiluminescent nitrogen analysis (Antek Instruments, Houston, Tex.)using caffeine as a standard. This data was utilized to determinedilution concentrations for subsequent spectral analysis. The extinctioncoefficients (ε) for LNO₂ and the isotopic derivatives [¹³C]LNO₂ and[¹⁵N]LNO₂ were measured using a UV-VIS spectrophotometer (Shimadzu,Japan) set to measure absorbance at 329 nm, the absorbance maximumspecific to LNO₂ as compared to linoleic acid. Absorbance values forincreasing concentrations of LNO₂, [¹⁵N]LNO₂ or [¹³C]LNO₂ in MeOHcontaining 20 mM NaOH were plotted against concentration to calculateslope/extinction coefficient.

Nitrated linoleic acid displays a characteristic absorption profile andmaximum, permitting determination of an extinction coefficient andproviding a facile method for measuring concentrations of syntheticLNO₂. This species displays a unique absorbance maximum at 329 nm,compared with linoleic acid (FIG. 3A). Plotting absorbance versusconcentration profiles for each of the synthetic LNO₂ preparationsreported herein—[¹⁴N]LNO₂, [¹⁵N]LNO₂ and [¹³C]LNO₂—generated identicalextinction coefficients for all LNO₂ derivatives: ε=10.1 cm⁻¹ M⁻¹ (FIG.3B).

Red Blood Cell and Plasma Lipid Isolation and Extraction.

Peripheral blood from healthy human volunteers was collected byvenipuncture in heparinized tubes, centrifuged (1200×g; 10 min) andplasma isolated from red cell pellets from which the buffy coat wasremoved. Crude lipid extracts were prepared from packed red cells andplasma by the method of Bligh and Dyer (29) and analyzed by massspectrometry. Care was taken to avoid acidification during all steps ofplasma fractionation and lipid extraction to prevent artifactual lipidnitration due to the presence of endogenous NO₂ ⁻. Extracts from redcells were analyzed by mass spectrometry; however, lipid extracts fromplasma were first fractionated by TLC to separate LNO₂ from the bulk ofneutral lipids present in plasma, minimizing ionization dampening duringLNO₂ analysis by mass spectrometry. TLC plates were developed twice in asolvent system consisting of hexane:ether:acetic acid (70:30:1, v/v),and regions of silica containing LNO₂, identified by comparing to themigration of synthetic standards, were scraped and extracted (29). Tomeasure the esterified LNO₂ content in red cell membranes and plasmalipoproteins, lipid extracts were first hydrolyzed (30), fractionated byTLC and analyzed by mass spectrometry.

In FIG. 6A, [¹³C]LNO₂ was added to lipid extractions as an internalstandard to quantitate the free and esterified LNO₂ content of red cellsand plasma obtained from healthy human volunteers. The mean age of the 5female and 5 male subjects was 34 yr. The endogenous LNO₂ isomersco-eluted with the added ¹³C-labeled LNO₂ internal standard, with theinternal standard differentiated from endogenous LNO₂ by monitoring itsunique m/z 342/295 MRM transition. In FIG. 6B, the internal standardcurve for LNO₂ reveals linear detector responses over five orders ofmagnitude. The limit of quantitation (LOQ) for LNO₂, as defined by tentimes the standard deviation of the noise, is approximately 0.3 fmol(˜100 fg) injected on column.

Analysis of Synthetic LNO₂ Methyl and Pentafluorobenzyl Esters by GasChromatography Mass Spectrometry.

Methyl ester-derivatives of synthetic LNO₂ isomers were analyzed by gaschromatography mass spectrometry (GC-MS) in both positive and negativeion modes. Electron-impact (EI) ionization was used to identify andcharacterize the fragmentation pattern of the two main positionalisomers of LNO₂ methyl esters. Methyl esters were prepared by dryingLNO₂ under a stream of nitrogen and redissolving in methanolic BF₃ (14%BF₃, 86% CH₃OH); with the methylation reaction proceeding for 8 min at60° C. Methyl esters were then extracted with hexane, washed twice withsaturated saline, redissolved in undecane and analyzed byelectron-impact GC-MS. EI GC-MS was performed using a Saturn 2000 massspectrometer coupled with a Varian 3800 gas chromatograph. Samples wereionized by electron impact at +70 eV. Methyl ester-derivatized LNO₂isomers were resolved by GC using a CP-7420 capillary column (0.25 mmID, 100 m fused silica, Varian, Palo Alto, Calif.) with the followingtemperature gradient: 60° C. (2 min); 60 to 120° C. at 20° C./min, heldfor 2 min; 120 to 270° C. at 20° C./min, held for 20 min. Helium wasused as the carrier gas.

Due to the low sensitivity of positive ion GC-MS to LNO₂, negative ionchemical ionization (NICI) was used to characterize pentafluorobenzyl(PFB) esters of synthetic LNO₂ and detect LNO₂ species in vivo. PFBesters of synthetic LNO₂, red cell and plasma lipids were prepared (29),with biological lipids first partially purified by TLC as previouslynoted. Lipids were then subjected to PFB esterification and analysis bynegative ion chemical ionization GC-MS using a Hewlett Packard 5890 GC(Palo Alto, Calif.) coupled to a single quadrupole Hewlett Packard MSusing a 30 m CP-Sil 8CB-MS column (5% phenyl, 95% dimethylpolysiloxane;Varian, Palo Alto, Calif.) (31). The following temperature gradient wasused to resolve the positional isomers of LNO₂: 165 to 190° C. at 10°C./min; 190 to 270° C. at 2° C./min; and 270 to 290° C. at 5° C./min.Helium and methane were used as carrier and reagent gases, respectively.The detection of LNO₂ was conducted by total ion count monitoring of[M-PFB]⁻, i.e., m/z 324 ([¹²C¹⁴N]LNO₂), 325 ([¹²C, ¹⁵N]) and 342([¹³C,¹⁴N]LNO₂).

Initial characterization of positional isomers of synthetic LNO₂ was viaEI GC-MS on LNO₂ methyl esters. Total ion count monitoring of thederivatized parent ion LNO₂ revealed 2 dominant peaks at 38.75 and 39min when resolved using a 100 m column (FIG. 4A). Product ion analysisof each peak (FIG. 4C) generated fragmentation patterns similar topreviously reported for acidic nitration of ethyl linoleate (18). Thefirst and second peaks correspond to linoleic acid that is nitrated onthe 12- and the 10-carbon, respectively (referred to as C12 and C10isomers). These two isomers are identified by the unique daughter ionsm/z 250 and m/z 282 (specific to C12), and m/z 196 (specific to C10).Analysis of [¹⁵N]LNO₂ revealed fragmentation patterns with theseidentifying ions shifted by m/z=+1, further affirming fragmentscontaining the nitro group (FIG. 4C). As a control for potentialBF₃-dependent artifactual products, anhydrous methanolic sulfuric acid(1%) esterification was performed to further verify the formation of thesame respective LNO₂ methyl ester positional isomers.

Structural analysis of synthetic LNO₂ isomers by EI GC MS yieldedimportant isomeric structural information and identification; but lackssensitivity for the detection, structural characterization andquantification of LNO₂ derivatives present in biological samples. Thus,these analyses were performed by NICI GC-MS on LNO₂ species that hadbeen derivatized to PFB esters (FIG. 4B). Chromatographic separation ofboth synthetic LNO₂ and TLC-separated red cell lipid extracts showed twodominant peaks, C12 and C10, and two minor peaks ascribed to C13 and C9positional isomers of LNO₂; with approximately 95% of total peak areaaccounted for by C12 and C10 isomers. To confirm that the identities ofthe LNO₂—PFB derivatives were the same as those identified asLNO₂-methyl esters, the LNO₂—PFB derivatives were run on EI positiveionization mode and identified by formation of characteristicfragmentation pattern (data not shown). Initial detection andquantitation of endogenous levels of LNO₂ was performed using NICIGC-MS. [¹³C]LNO₂ was added during the monophase of lipid extractions asan internal standard to correct for losses due to TLC andderivatization. The requirement for TLC separation to enrich fractionscontaining LNO₂, the additional requirement for derivatization andon-column thermal decomposition creating limitations for analyzingnitrohydroxy- and nitrohydroperoxy fatty acids all combined to yieldinconsistent quantitation of LNO₂ levels in biological samples uponanalysis by GC-MS. Consequently, we developed a LC-MS/MS based method tomore reliably characterize and quantitate LNO₂ in biological samples.

Analysis of LNO₂ Positional Isomers by Electrospray Ionization TripleQuadrupole Mass Spectrometry.

Qualitative analysis of nitrated linoleic acid positional isomers byelectrospray ionization mass spectrometry was performed using an AppliedBiosystems/MDS Sciex 4000 Q Trap™, a hybrid triple quadrupole-linear iontrap mass spectrometer. To separate and characterize the two major LNO₂positional isomers, synthetic standards and lipid extracts frombiological samples were resolved by reverse-phase high performanceliquid chromatography (HPLC) using a 150×2 mm C18 Phenomenex Luna column(3 μm particle size). Lipids were eluted using a gradient solvent systemconsisting of A (H₂O containing 0.1% NH₄OH) and B (CNCH₃ containing 0.1%H₂O) under the following conditions: 20-70% B (linear increase, 20 min),70-95% B (2 min), 95% B (8 min), 95-20% B (1 min) and 20% B (5 min).Using these gradient conditions, two major and two minor LNO₂ positionalisomers were separated with baseline resolution. The resolved positionalisomers of LNO₂ were detected by mass spectrometry using a multiplereaction monitoring (MRM) scan mode by reporting molecules that undergoan m/z 324/277 mass transition. This transition, consistent with theloss of HNO₂ ([M-(HNO₂)—H]⁻), is common for all mono-nitrated positionalisomers of linoleic acid. Concurrent with MRM, enhanced product ionanalysis (EPI) was performed to generate characteristic and identifyingfragmentation patterns of the eluting species with a precursor mass ofm/z 324. In some analyses, specific fragments appearing in the EPIspectra were further fragmented in the ion trap to generate a MS³spectrum to verify structural elucidation.

The most sensitive and selective technique available for analytequantitation is triple quadrupole mass spectrometry detecting in theMRM. An HPLC separation strategy that baseline-resolved individual LNO₂positional isomers permitted the MS-based quantitation of LNO₂positional isomers, following chromatography of red cell and plasmalipid extracts. By monitoring the m/z 324/277 mass transition of thelinoleate nitro derivative and the fatty acid parent molecule,respectively (FIG. 5A-1). To initially identify individual LNO₂positional isomers resolved by HPLC separation, the two major specieswere collected (peaks 1 and 2, FIG. 6B), derivatized to PFB-esters andanalyzed by GC-MS. From GC-MS analysis, the first species eluting uponHPLC separation was identified as the C12 isomer and the second the C10nitro derivative. To further characterize synthetic LNO₂ isomericstructures, the linear ion trap mode of the hybrid mass spectrometer wasused to perform enhanced product ion analysis on eluting peaks togenerate a characteristic, identifying fragmentation pattern for eachLNO₂ positional isomer (FIGS. 5B-1, C-1). The ion trap enabled theconcentration and thus detection of otherwise minor fragment ions, withMS/MS analysis revealing the unique fragments m/z 168 and 228 for theC10 isomer, as well as the fragments common to all LNO₂ isomers (m/z233, 244, 277, 293 and 306, FIG. 5C-1). The m/z 228 ion was furtherfragmented in the ion trap using the MS³ scanning mode, generating afragment with m/z 46, indicating the loss of a nitro group and assigningthe nitro group to the 9-10 double bond. Using [¹⁵N]LNO₂, the C10 isomergenerated a fragment of m/z 229 that undergoes further fragmentation tom/z 47 (not shown). FIG. 5B-1 presents the characteristic fragmentationpattern obtained from MS/MS of the C12 isomer. The unique fragments toC12-LNO₂, m/z 196 and 157, and the common fragments (m/z 233, 244, 277,293 and 306) are present and consistent with a nitro group located atthe 12-carbon of linoleic acid. In aggregate, from the syntheticrationale and GC MS and HPLC-ESI MS/MS data, the C10 and the C12 isomersare identified as 10-nitro-9-cis,12-cis-octadecadienoic acid and12-nitro-9-cis,12-cis-octadecadienoic acid, respectively.

HPLC-ESI MS/MS was used to characterize and quantitate LNO₂ speciespresent in healthy human blood, specifically red cells and plasma (FIG.5). The MRM elution profiles for red cell and plasma lipid extracts wereidentical to those obtained from the synthetic standards (FIGS. 5B-2,C-2 and 5B-3, C-3). All characteristic fragments found in the EPIspectra of the synthetic C10 and C12 LNO₂ isomers were present in theresolved biological extracts, further affirming that red cells andplasma contain LNO₂ positional isomers identical to synthetic standards.

Control Studies Relevant to Processing and Analysis-Induced LipidNitration.

In the presence of nitrite and acidic conditions, unsaturated fattyacids undergo nitration to products that reflect the chromatographic andMS characteristics of nitrosenylation-derived synthetic LNO₂ isomers(18-20). Thus, acidic conditions were either avoided or documented forpotential influence on product yield and nature. Importantly, lipidextractions were routinely conducted in the presence of pH 7.4 aqueousbuffers. Comparison of LNO₂ content in preparations extracted underneutral or acidic conditions revealed that there was no difference inLNO₂ yield or LNO₂ isomer distribution only in the complete absence ofNO₂ ⁻. Acidic lipid extraction conditions (pH<4.0) in the presence ofexogenously-added biological concentrations of NO₂ ⁻ (10-500 μM) inducedadditional linoleate nitration, as indicated by nitration of¹³C-linoleic acid added to pure linoleate or biological lipid extracts.When lipid extracts are separated by TLC prior to MS analysis, effectiveresolution of LNO₂ from native and oxidized fatty acids requires the useof a 1% acetic acid-containing solvent system. It was observed that whenNO₂ ⁻ was present in the TLC chromatographic solvent at concentrationsnot expected from biological analyses (>1 mM), LNO₂ was formed de novoby acid-catalyzed nitration reactions. Since the initial solventextraction of lipids from synthetic mixtures or biological materialsresulted in removal of >95% of all adventitious NO₂ ⁻, confidence existsthat artifactual fatty acid nitration reactions were not occurringduring acidic HPLC or TLC resolution. With regard to biological sampleanalysis, this latter precept was affirmed by a) adding ¹³C-labeledlinoleic acid prior to red cell and plasma lipid extraction,purification and MS analysis and b) observing no formation of¹³C-labeled nitrated linoleic acid derivatives.

Detection and Quantitation of LNO₂ in Human Red Blood Cells.

Quantitation of LNO₂ in biological samples was performed as above, withmodification. During the monophase stage of lipid extractions (29), aknown quantity of [¹³C]LNO₂ was added as internal standard to correctfor loss during extraction and TLC. The gradient elution profile waschanged so that all LNO₂ positional isomers co-eluted. Lipids wereeluted from the same column used for structural characterization using agradient solvent system consisting of A (H₂O containing 0.1% NH₄OH) andB (CNCH₃ containing 0.1% H₂O) under the following conditions: 80-90% B(2 min), 90% B (3 min), 90-80% B (1 min) and 80% B (2 min). Two MRMtransitions were monitored: m/z 324/277, for LNO₂ isomers, and m/z342/295, a transition consistent with the loss of HNO₂ for the¹³C-labeled internal standard. The areas under each peak wereintegrated, the ratio of analyte to internal standard areas wasdetermined and amounts of LNO₂ were quantitated using Analyst 1.4quantitation software (Applied Biosystems, Foster City, Calif.) byfitting the data to an internal standard curve. Adjustable massspectrometer settings for both qualitative and quantitative analyseswere as follows: CUR 10; IS −4500; TEM 450; GS1 50; GS2 60; CAD 2 (8when EPI experiments were concurrently performed); DP −50; and EP −10.House-generated zero grade air was used as the source gas and nitrogenwas used as the curtain and collision gases.

To quantitate net LNO₂ species present in red cells and plasma, HPLCgradient conditions were changed (see Experimental Methods) so that allpositional isomers co-eluted (FIG. 6). MRM transitions for the combinedLNO₂ and [¹³C]LNO₂ (m/z 324 and 342, respectively) were monitored, withanalytes eluting at 2 min. Nitrated linoleic acid concentration was afunction of the ratio of analyte to internal standard peak areas usingan internal standard curve that is linear over five orders of magnitude.Blood samples obtained from ten healthy human volunteers (5 female, 5male, ages ranging from 22 to 45) revealed free LNO₂ in red cells (i.e.,LNO₂ not esterified to glycerophospholipids or neutral lipids) to be49.6±16.6 pmol/ml packed cells. Total free and esterified LNO₂, theamount present in saponified samples, was 249±104 pmol/ml packed cells.Thus, approximately 75% of LNO₂ in red cells exists as esterified fattyacids. In plasma, free and total (free plus esterified) LNO₂ was78.9±34.5 pmol/ml plasma 629±242 nM, respectively, with free LNO₂representing 85% of total (Table 1). These values indicate that LNO₂species are quantitatively the greatest pool of bioactive oxides ofnitrogen in the vascular compartment (34-37).

Throughout the Background and Example 1, several publications have beenreferenced. These publications are listed in the Reference Listfollowing Example 1. The disclosures of these publications in theirentireties are hereby incorporated by reference into this application inorder to more fully describe the compounds, compositions and methodsdescribed herein.

TABLE 1 Biologically active nitrogen oxide derivatives in human blood -comparison with net C10- and C12-nitro derivatives of linoleic acid.Venous blood was obtained from healthy human volunteers, centrifuged(1200 x g; 10 min) and plasma isolated from red cell pellets from whichthe buffy coat was removed. Total lipid extracts were prepared frompacked red cells and plasma (29) and analyzed by mass spectrometry asdescribed in Experimental Methods. Total LNO₂ (free plus esterified) wasdetermined in lipid extracts following saponification. Free and totalLNO₂ was quantitated by fitting analyte to internal standard area ratiosobtained by mass spectrometry to an internal standard curve. Data areexpressed as mean ± SD (n = 10; 5 female, 5 male). Species CompartmentConcentration (nM) Reference NO₂ ⁻ Plasma 205 ± 21  (35, 36) RSNO Plasma7.2 ± 1.1 (35, 36) Plasma LNO₂ Free 80 ± 34 Esterified 550 ± 274 Total630 ± 240 Hb-NO Blood >50 (37) Hb-SNO Blood 0-150 (34) Packed red cellsLNO₂ Free 25 ± 8  Esterified 199 ± 60  Total 224 ± 52  LNO₂ Whole Blood*477 ± 128 *assuming a 40% hematocrit

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Example 2

Allylic nitro derivatives of linoleic acid (nitrolinoleic acid, LNO₂)are formed via nitric oxide-dependent oxidative inflammatory reactionsand are found at concentrations of ˜500 nM in the blood of healthyindividuals. It will be shown that nitrolinoleic acid (LNO₂) is a potentendogenous ligand for peroxisome proliferator-activated receptor γ (PPARγ; K_(i)˜133 nM) that acts within physiological concentration ranges.This nuclear hormone receptor (PPARγ) regulates glucose homeostasis,lipid metabolism and inflammation. PPARγ ligand activity is specific forLNO₂ and not mediated by LNO₂ decay products, NO donors, linoleic acidor oxidized linoleic acid. LNO₂ is a significantly more robust PPARγligand than other reported endogenous PPARγ ligands, includinglysophosphatidic acid (LPA 16:0 and LPA 18:1), 15-deoxy-Δ^(12,14)-PGJ₂,conjugated linoleic acid and azelaoyl-PC. LNO₂ activation of PPAR γ viaCV-1 cell luciferase reporter gene expression analysis revealed a ligandactivity that rivals or exceeds synthetic PPARγ agonists such asRosiglitazone and Ciglitazone, is co-activated by 9 cis-retinoic acidand is inhibited by the PPAR γ antagonist GW9662. LNO₂ inducesPPARγ-dependent macrophage CD-36 expression, adipocyte differentiationand glucose uptake also at a potency rivaling thiazolidinediones. Theseobservations reveal that nitric oxide (.NO)-mediated cell signalingreactions can be transduced by fatty acid nitration products andPPAR-dependent gene expression.

The reaction of nitric oxide (.NO) with tissue free radical andoxidative intermediates yields secondary oxides of nitrogen that mediateoxidation, nitration and nitrosation reactions (1, 2). Of presentrelevance, the reaction of .NO and .NO-derived species with oxidizingunsaturated fatty acids is kinetically rapid and exerts a multifacetedimpact on cell redox and signaling reactions. Nitric oxide readilyout-competes lipophilic antioxidants for the scavenging of lipidradicals, resulting in the inhibition of peroxyl radical-mediated chainpropagation reactions (3). Both the catalytic activity and geneexpression of eicosanoid biosynthetic enzymes are also regulated by .NO,affirming a strong linkage between .NO and fatty acid oxygenationproduct synthesis and signaling (4, 5). Consistent with this latterprecept, fatty acid nitration products generated by .NO-derived speciesinhibit multiple aspects of inflammatory cell function, indicating thatnitrated fatty acids are both byproducts and mediators of redoxsignaling reactions (6-8).

Recently, the structural characterization and quantitation ofnitrolinoleic acid (LNO₂) in human red cells and plasma revealed thisunsaturated fatty acid derivative to be the most abundant bioactiveoxide of nitrogen in the vasculature. Net blood levels of ˜80 and 550 nMfree and esterified LNO₂, respectively, were measured in healthy humans(9). The observation that .NO-dependent oxidative inflammatory reactionsyields allylic nitro derivatives of unsaturated fatty acids displayingcGMP-independent cell signaling properties (5) led to the identificationof a receptor that can transduce LNO₂ signaling. Affymetrixoligonucleotide microarray analysis of cRNA prepared from methanol(vehicle), linoleic acid (LA)- and LNO₂-treated human aortic smoothmuscle cells indicated that LNO₂ specifically and potently regulated theexpression of key inflammatory, cell proliferation and celldifferentiation-related proteins. Multiple PPARγ target genes weresignificantly regulated, suggesting that LNO₂ serves as an endogenousPPARγ ligand.

PPARγ is a nuclear hormone receptor that binds lipophilic ligands.Downstream effects of PPARγ activation include modulation of metabolicand cellular differentiation genes and regulation of inflammatoryresponses (e.g., integrin expression, lipid transport by monocytes),adipogenesis and glucose homeostasis (10, 11). In the vasculature, PPARγis expressed in monocytes, macrophages, smooth muscle cells andendothelium (12) and plays a central role in regulating the expressionof genes related to lipid trafficking, cell proliferation andinflammatory signaling(13). While synthetic thiazolidinediones (TZDs)such as Rosiglitazone and Ciglitazone are appreciated to be the mostpotent PPARγ ligands yet described, considerable interest and debateremains focused on the identity of endogenous PPARγ ligands because oftherapeutic potential and their intrinsic value in understanding cellsignaling. At present, tissue and plasma levels of putative PPAR ligandsare frequently not precisely defined and when so, are found inconcentrations sometimes orders of magnitude lower than those requiredto activate specific α, γ or δ PPAR subtypes (14-16). As set forthherein, the allylic nitro derivatives of fatty acids are robustendogenous PPARγ ligands that act within physiological concentrationranges to modulate key PPARγ-regulated signaling events includingadipogenesis, adipocyte glucose homeostasis and CD36 expression inmacrophages.

Materials.

LNO₂ and [¹³C]LNO₂ were synthesized and purified using linoleic acid(NuCheckPrep, Elysian, Minn.) and [¹³C]linoleic acid (Spectra StablesIsotopes, Columbia, Md.) subjected to nitroselenylation as previouslydescribed (9). LNO₂ concentrations were quantified spectroscopically andby chemiluminescent nitrogen analysis (Antec Instruments, Houston, Tex.)using caffeine as a standard (9). Anti-CD36 antibody (kindly provided byDr. de Beer at University of Kentucky Medical Center, Lexington, Ky.);anti-PPARγ and anti-β-actin antibodies were from Santa Cruz (Santa Cruz,Calif.); and anti-aP2 antibody was from Chemicon International Inc.(Temecula, Calif.). Horseradish peroxidase-linked goat anti-rabbit IgGand Coomasie Blue were from Pierce (Rockford, Ill.). [³H]Rosiglitazonewas from American Radiolabeled Chemical, Inc. (St. Louis, Mo.). [³H]2-Deoxy-D-glucose was from Sigma (St Louis, Mich.). Scintil-safe plus TM50% was from Fisher Scientific (Pittsburgh, Pa.). Rosiglitazone,Ciglitazone, 15-deoxy-Δ^(12,14)-PGJ₂, Conjugated Linoleic Acid (CLA1,CLA2) and GW9662 were from Cayman Chemical (Ann Arbor, Mich.).1-palmitoyl-2-hydroxy-sn-glycero-3-Phosphate (16:0 LPA),1-O-9-(Z)-octadecenyl-2-hydroxy-sn-glycero-3-phosphate (18:1 LPA),1-O-hexadecyl-2-azelaoyl-sn-glycero-3-phosphocholine (azPC),1-Palmitoy-2-Azelaoyl-sn-Glycero-3-Phosphocholine (azPC Ester) were fromAvanti Polar Lipids, Inc. (Alabaster, Ala.).

Cell Transient Transfection Assay.

CV-1 cells from ATCC (Manassas, Va.) were grown to ˜85% confluence inDMEM/F12 supplemented with 10% FBS, 1% penicillin-streptomycin. Then,cells were transiently co-transfected with a plasmid containing theluciferase gene under regulation by four Gal4 DNA binding elements(UAS_(G)×4 TK-Luciferase, a gift from Dr. Ronald M. Evans), in concertwith plasmids containing the ligand binding domain for the differentnuclear receptors fused to the Gal4 DNA binding domain. For assessingfull-length PPAR receptors, CV-1 cells were transiently co-transfectedwith a plasmid containing the luciferase gene under the control of threetandem PPAR response elements (PPRE) (PPRE×3 TK-Luciferase) and hPPARγ,hPPARα or hPPARδ expression plasmids, respectively. In all cases,fluorescence protein (GFP) expression plasmid was co-transfected as thecontrol for the transfection efficiency. Twenty-four hours after thetransfection, cells were cultured for another 24 hours in Optium-MEM(Invitrogen, Carlsbad, Calif.). Then, cells were treated with differentcompounds as indicated in figures for 24 hours in Optium-MEM. Reporterluciferase assay kits from Promega (Madison, Wis.) were used to measurethe luciferase activity according to the manufacturer's instructionswith a luminometer (Victor II, Perkin-Elmer). Luciferase activity wasnormalized by GFP units. Each condition was performed at least intriplicates in each experiment. All experiments were repeated at leastthree times.

PPARγ Competition Binding Assay.

Human PPARγ1 cDNA was inserted into pGEX from Amersham Biosciences Corp(Piscataway, N.J.) containing the gene encoding glutathioneS-transferase (GST). GST-PPARγ protein induction and receptor bindingwas assessed as previously described (17).

3T3-L1 Differentiation and Oil Red 0 Staining.

3T3-L1 preadipocytes were propagated and maintained in DMEM containing10% FBS. To induce differentiation, 2-day postconfluent preadipocytes(designated day 0) were cultured in DMEM containing 10% FBS plus 3 μMLNO₂ for 14 days. The medium was changed every two days. Rosiglitazone(3 μM) and linoleic acid (10 μM) were used as the positive and negativecontrol, respectively. The differentiated adipocytes were stained by Oilred O as previously described (18).

[³H]-2-Deoxy-D-glucose Uptake Assay in Differentiated 3T3-L1 Adipocyte.

[³H]-2-Deoxy-D-glucose uptake assay as previously described (19). 3T3-L1preadipocytes were grown in 24-well tissue culture plates, 2-daypostconfluent preadipocytes were treated by 10 μg/ml insulin (Sigma), 1μM dexamethasone (Sigma), and 0.5 mM 3-isobutyl-1-methylxanthine (Sigma)in DMEM containing 10% FBS for two days, then cells were kept in 10μg/ml insulin also in DMEM containing 10% FBS for 6 days (changed mediumevery three days). Eight days after induction of adipogenesis, testcompounds in DMEM containing 10% FBS were added for an additional 2 days(changed medium every day) and PPARγ-specific antagonist GW9662 werepretreated 1 h before other additions. After two rinses with serum-freeDMEM, cells were incubated for 3 h in serum-free DMEM and rinsed at roomtemperature three times with freshly prepared KRPH buffer (5 mMphosphate buffer, 20 mM HEPES, 1 mM MgSO₄, 1 mM CaCl₂, 136 mM NaCl, 4.7mM KCl, pH 7.4). The buffer was replaced with 1 μCi/ml of[³H]-2-deoxy-D-glucose in KRPH buffer for 10 min at room temperature.The treated cells were rinsed carefully three times with cold PBS, lysedovernight in 0.8N NaOH (0.4 ml/well), and neutralized with 13.3 μl of12N HCl. Lysate (360 μl) was added to 4 ml Scinti-safe plus TM 50% in ascintillation vial, and the vials were mixed and counted.

RNA and Protein Preparation and Analysis.

RNA and protein expression levels were analyzed by quantitativereal-time PCR and Western blot analysis as previously described.²⁵

LNO₂ Decay.

LNO₂ decay was induced by incubating 3 μM LNO₂ in medium+serum at 37° C.At different times, samples were removed and analyzed for bioactivity orfor LNO₂ content by Bligh and Dyer extraction in the presence of 1 μM[¹³C]LNO₂ as an internal standard. Non-decayed LNO₂ was quantified viatriple quadrupole mass spectrometric analysis (Applied Biosystems/MDSSciex, Thornhill, Ontario, Canada) as previously reported (9).

To characterize LNO₂ as a potential ligand for a lipid-binding nuclearreceptor [e.g., PPARα, PPARγ, PPARγ, androgen receptor (AR),glucocorticoid receptor (GR), mineralocorticoid receptor (MR),progesterone receptor (PR) and retinoic X receptor α (RXRα)], CV-1reporter cells were co-transfected with plasmids containing the ligandbinding domain for these nuclear receptors fused to the Gal4 DNA bindingdomain and the luciferase gene under regulation of four Gal4 DNA bindingelements. LNO₂ (1 μM) induced significant activation of PPARγ (620%),PPARα (325%) and PPARγ (221%), with no impact on AR, GR, MR, PR or RXRαreceptor activation (FIG. 7A). To further explore PPAR activation byLNO₂, CV-1 cells were transiently co-transfected with a plasmidcontaining the luciferase gene under three PPAR response elements (PPRE)in concert with PPARγ, PPARα or PPARγ expression plasmids.Dose-dependent activation by LNO₂ was observed for all PPARs by LNO₂(FIG. 7B), with PPARγ showing the greatest response atclinically-relevant concentrations of LNO₂.

PPARγ activation by LNO₂ rivaled that induced by Ciglitazone andRosiglitazone and exceeded that of 15-deoxy-Δ^(12,14)-PGJ₂(20, 21),which only occurred at concentrations 3 orders of magnitude greater thanfound clinically (FIG. 7C and inset). Several other reported endogenousPPARγ activators added in equimolar concentration with LNO₂ [linoleicacid (LA), conjugated linoleic acid (CLA1, CLA2), lysophosphatidic acid(LPA-16:0, LPA-18:1), azPC (azelaoylPC) and azPC ester; 1 and 3 μM],displayed no significant activation of PPARγ reporter gene expressionwhen compared with vehicle control (FIG. 7C) (22-24) LNO₂-mediated PPARγactivation was inhibited by the PPARγ-specific antagonist GW9662 andenhanced ˜180% by co-addition of the RXRα agonist 9-cis-retinoic acid,which facilitates PPRE promoter activation via heterodimerization ofactivated RXRα with PPARγ (FIG. 8A).

LNO₂ slowly undergoes decay reactions in aqueous solution, displaying a30-60 min half-life (FIG. 8B) and yielding .NO and an array of oxidationproducts. The activation of PPARγ paralleled the presence andconcentration of the LNO₂ parent molecule, as determined by concomitantreceptor activation analysis and mass spectrometric quantitation ofresidual LNO₂ in reaction mixtures undergoing different degrees ofaqueous decomposition prior to addition to CV-1 cells (FIG. 8B).Affirmation that activation of PPARγ is LNO₂-specific, rather than aconsequence of LNO₂ decay products, was established by treatment ofPPARγ-transfected CV-1 cells with .NO donors and oxidized linoleic acidderivatives. Additionally, the fatty acid oxidation products 9- and13-oxoODE are reported endogenous PPARγ stimuli (25-27). 13-oxoODE hadno effect on PPARγ-dependent reporter gene expression (FIG. 8C). Onlyhigh and non-physiological concentrations of 9-oxoODE, and the concertedaddition of high concentrations of 13-oxoODE and 5-nitrosoglutathione orspermine-NONOate, resulted in modest PPARγ activation (FIG. 8C). The .NOdonors added individually did not activate PPARγ, even at highconcentrations, eliminating the possibility that PPARγ-mediatedsignaling by LNO₂ is due to .NO derivatives released during LNO₂ decay(FIG. 8C).

Competitive PPARγ binding analysis quantified the displacement of[³H]Rosiglitazone by unlabeled Rosiglitazone, LNO₂ and linoleic acid.The calculated binding affinity (Ki) for Rosiglitazone was 53 nM,consistent with reported values (FIG. 8D, 40-50 nM (28)). LNO₂ displayedan estimated Ki of 133 nM and linoleic acid an estimated Ki of >1,000 nM(previously reported as 1.7 to 17 μM (28)). This reveals that the Ki forLNO₂ displacement of [³H]Rosiglitazone from PPARγ is comparable to thishighly avid ligand.

The PPARγ agonist actions of LNO₂ were also examined in a biologicalcontext, using cell models noted for well-established PPARγ-dependentfunctions. The scavenger receptor CD36 is expressed in diverse celltypes, including platelets, adipocytes and macrophages. In macrophages,CD36 is a receptor for oxidized LDL, with expression positivelyregulated by PPARγ (29). Treatment of mouse RAW264.7 macrophages withLNO₂ induced greater CD36 receptor protein expression than an equivalentRosiglitazone concentration, a response partially inhibitable by thePPARγ-specific antagonist GW9662 (FIG. 9A). Moreover, quantitativereal-time PCR revealed that LNO₂-dependent PPARγ transactivation induceda dose-dependent increase in CD36 mRNA expression in these macrophages.

PPARγ plays an essential role in the differentiation of adipocytes (30,31). In support of this precept, selective disruption of PPARγ resultsin impaired development of adipose tissue (18, 32). To define if LNO₂induces PPARγ-activated adipogenesis, 3T3-L1 preadipocytes were treatedwith LNO₂, Rosiglitazone or linoleic acid for two weeks. Adipocytedifferentiation was assessed both morphologically and via oil red Ostaining, which reveals the accumulation of intracellular lipids.Vehicle and linoleic acid did not affect differentiation, while LNO₂induced >30% of 3T3-L1 preadipocyte differentiation (FIG. 9B).Rosiglitazone treatment affirmed a positive PPARγ-dependent response.LNO₂ and Rosiglitazone-induced preadipocyte differentiation alsoresulted in expression of specific adipocyte markers (PPARγ2 and aP2),an event not detected for linoleic acid (FIG. 9C). PPARγ ligands play acentral role in glucose metabolism, with TZDs widely used asinsulin-sensitizing drugs. Addition of LNO₂ (1-10 μM) to differentiatedadipocytes induced a dose-dependent increase in glucose uptake (FIG. 9D,Left panel). The impact of LNO₂ on glucose uptake was greater than thatobserved for equimolar 15-deoxy-PGJ₂, equivalent to Rosiglitazone andsimilarly inhibited by the PPARγ-specific antagonist GW9662 (FIG. 9D,Right panel). In aggregate, these observations establish that LNO₂induces well-characterized PPARγ-dependent signaling actions towardsmacrophage CD36 expression and adipogenesis.

The identification of bona fide high affinity endogenous PPARγ ligandshas been a provocative issue that, when resolved, will advance ourunderstanding of endogenous PPARγ modulation and can reveal new meansfor intervention in diverse metabolic disorders and disease processes.This will also shed light on the broader contributions of this nuclearhormone receptor family to both the maintenance of tissue homeostasisand the regulation of cell and organ dysfunction. Of relevance, thegeneration of PPAR-activating intermediates from complex lipids oftenrequires hydrolysis by phospholipase A₂ (PLA₂), with the identity ofphospholipid sn-2 position fatty acid derivatives that might serve asPPAR agonists still forthcoming (33, 34). Thus, the fact that ˜80% ofLNO₂ present in a variety of tissue compartments is esterifiedencourages the notion that this pool of high affinity PPAR ligandactivity will be mobilized upon the PLA₂ activation that occurs duringinflammation to regulate the formation of cell signaling molecules.

Comparison of PPARγ-dependent gene expression induced by LNO₂ with thatof TZDs and putative fatty acid- and phospholipid-derived PPARγ ligandsfurther affirmed the robust activity of LNO₂ as a PPARγ ligand. While anumber of endogenous lipophilic species are proposed as PPARγ ligands,their intrinsically low binding affinities and in vivo concentrations donot support a capability to serve as physiologically-relevant signalingmediators. Presently-reported endogenous PPARγ agonists include freefatty acids, components of oxidized plasma lipoproteins (9- and13-oxoODE, azPC), conjugated linoleic acid derivatives (CLA1 and CLA2),products of phospholipase hydrolysis of complex lipids (LPA), plateletactivating factor (PAF) and eicosanoid derivatives such as thedehydration product of PGD₂, 15-deoxy-Δ^(12,14)-PGJ₂ (14, 22, 23).Herein, minimal or no activation of PPARγ reporter gene expression by1-3 μM concentrations of these putative ligands was observed, incontrast to the dose-dependent PPAR activation by LNO₂ that, for PPARγ,was significant at clinically-relevant concentrations as low as 100 nM.A dilemma exists in that some putative endogenous PPARγ agonists haveonly been generated by aggressive in vitro oxidizing conditions (e.g.,Cu-mediated LDL oxidation) and have not been clinically quantified ordetected (for instance, azPC, CLA). Other lipid derivatives proposed asPPAR ligands are present in <100 nM tissue concentrations, orders ofmagnitude below their binding affinities (1-15 μM) and are not expectedto result in significant receptor occupancy and activation in vivo. Thislatter category includes free fatty acids, eicosanoids, 9- and13-oxoODE, PAF and 15-deoxy-Δ^(12,14)-PGJ₂ (14). For example, while LPAis a notable PPARγ ligand of relevance to vascular, inflammatory andcell proliferative diseases the plasma concentration of LPA is wellbelow 100 nM (35, 36) and its binding affinity for PPARγ has not beenestablished. Also, the in vivo downstream vascular signaling actions ofLPA are inconsistent with its proposed PPARγ ligand activity (24,37-39). Future studies using mice genetically deficient for PPARγ andNOS isoforms should assist in defining the mechanisms of formation andendogenous PPAR ligand activity of nitrated fatty acids.

In summary, LNO₂ is a high affinity ligand for PPARs, especially PPARγ,that activates both reporter constructs and cells at physiologicalconcentrations. Fatty acid nitration products, generated by.NO-dependent reactions, are thus expected to display broad cellsignaling capabilities as endogenous nuclear receptor-dependentparacrine signaling molecules with a potency that rivals TZDs (FIG. 7C).Present data reveals that LNO₂ mediates cell differentiation inadipocytes, CD36 expression in macrophages, and inflammatory-relatedsignaling events in endothelium (e.g., inhibition of VCAM-1 expressionand function, not shown) with an important contribution fromPPARγ-dependent mechanisms (FIG. 9A-D). Allylic nitro derivatives offatty acids thus represent a unique class of receptor-dependent celldifferentiation, metabolic and anti-inflammatory signaling moleculesthat serve to converge .NO and oxygenated lipid redox signalingpathways.

Throughout Example 2, several publications have been referenced. Thesepublications are listed in the Reference List for Example 2. Thedisclosures of these publications in their entireties are herebyincorporated by reference into this application in order to more fullydescribe the compounds, compositions and methods described herein.

REFERENCE LIST FOR EXAMPLE 2

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V., Silva, A. R., St Hilaire, A.,    Xu, Y., Hinshaw, J. C., Zimmerman, G. A., Hama, K., Aoki, J.,    Arai, H. et al. (2003) Proc. Natl. Acad. Sci. U.S.A 100, 131-136.-   24. Zhang, C., Baker, D. L., Yasuda, S., Makarova, N., Balazs, L.,    Johnson, L. R., Marathe, G. K., McIntyre, T. M., Xu, Y.,    Prestwich, G. D. et al. (2004) J. Exp. Med. 199, 763-774.-   25. Kliewer, S. A., Sundseth, S. S., Jones, S. A., Brown, P. J.,    Wisely, G. B., Koble, C. S., Devchand, P., Wahli, W., Willson, T.    M., Lenhard, J. M. et al. (1997) Proc. Natl. Acad. Sci. U.S.A 94,    4318-4323.-   26. Nagy, L., Tontonoz, P., Alvarez, J. G., Chen, H. &    Evans, R. M. (1998) Cell 93, 229-240.-   27. Von Knethen, A. & Brune, B. (2002) J. Immunol. 169, 2619-2626.-   28. Ferry, G., Bruneau, V., Beauverger, P., Goussard, M., Rodriguez,    M., Lamamy, V., Dromaint, S., Canet, E., Galizzi, J. P. &    Boutin, J. A. (2001) Eur. J. Pharmacol. 417, 77-89.-   29. 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Example 3

The synthesis, structural characterization, clinical quantitation andcell signaling activity of nitrated oleic acid (OA-NO₂) are reported.Analysis of plasma and urine also revealed the presence of additionalnitrated fatty acids, including nitrated linolenic, arachidonic,eicosapentaenoic and docosahexaenoic acids and multiple nitrohydroxyderivatives, which reveal the ubiquity of nitrated fatty acidderivatives in humans. Two nitroalkene derivatives of oleic acid weresynthesized (9- and 10-nitro-9-cis-octadecenoic acid), structurallycharacterized and compared with nitrated fatty acids present in plasma,red cells and urine of healthy humans. Based on HPLC elution and massspectrometric characteristics, these two regioisomers of OA-NO₂ wereidentified in clinical samples. Using ¹³C isotope dilution, OA-NO₂ wasquantitated, with plasma free and esterified levels of 619±52 and302±369 nM, respectively. In red blood cells, free and esterified OA-NO₂was 59±11 and 155±65 nM, respectively. Assuming a 40% hematocrit, OA-NO₂levels are ˜50% greater than that of nitrated linoleic acid; withcombined free and esterified levels of these two nitroalkene derivativesexceeding 1 μM. OA-NO₂ potently activated peroxisome proliferatoractivated receptor-γ and induced PPARγ-dependent adipogenesis anddeoxyglucose uptake in 3T3 L1 preadipocytes. These data reveal thatmultiple nitrated fatty acids comprise a class of .NO and fattyacid-derived signaling molecules.

The oxidation of unsaturated fatty acids converts lipids, otherwiseserving as cell metabolic precursors and structural components, intopotent signaling molecules including prostaglandins, leukotrienes,isoprostanes, hydroxy- and hydroperoxy-eicosatetraenoates andplatelet-activating factor. This process, either enzymatic orauto-oxidative, orchestrates immune responses, neurotransmission and theregulation of cell growth. For example, prostaglandins arecyclooxygenase-derived lipid mediators that signal via receptor-ligandinteractions to regulate inflammatory responses, vascular function,initiation of parturition, cell survival and angiogenesis (1). Incontrast, the various isoprostane products of arachidonic acidauto-oxidation exert vasoconstrictive and pro-inflammatory signalingactions via both receptor-dependent and -independent mechanisms (2). Acommon element of these diverse lipid signaling actions is that nitricoxide (.NO) and other reactive nitrogen species significantly impactlipid mediator formation and bioactivities.

The ability of .NO and .NO-derived reactive species to oxidize,nitrosate and nitrate biomolecules suggests that .NO might alsoinfluence the synthesis and reactions of bioactive lipids (3-5).Interactions between .NO and lipid oxidation pathways are multifacetedand interdependent. For example, .NO regulates both the activity andexpression of prostaglandin H synthase (6). Conversely, leukotrieneproducts of lipoxygenases induce nitric oxide synthase-2 expression toincrease .NO production (7). Furthermore, the autocatalytic chainpropagation reactions of lipid peroxyl radicals during membrane andlipoprotein oxidation are potently inhibited by .NO (8). Of relevance,reactions between .NO-derived species and lipid oxidation intermediatesyield nitrated fatty acids. Recently, the nitroalkene derivative oflinoleic acid (LNO₂) has been detected in human blood at concentrationssufficient to induce biological responses (˜500 nM, refs. (9-12)).Compared with other .NO-derived species such as nitrite (NO₂),nitrosothiols (RSNO) and heme-nitrosyl complexes, LNO₂ represents thesingle most abundant pool of bioactive oxides of nitrogen in the healthyhuman vasculature (9, 13-16).

In vitro studies have shown that LNO₂ mediates cGMP-dependent vascularrelaxation, cGMP-independent inhibition of neutrophil degranulation andsuperoxide formation, and inhibition of platelet activation (10-12).Recently, LNO₂ has been shown to exert cell signaling actions vialigation and activation of peroxisome proliferator activated receptors(PPAR) (17), a class of nuclear hormone receptors that modulates theexpression of metabolic and cellular differentiation andinflammatory-related genes (18,19).

The identification of LNO₂ as an endogenous PPARγ ligand that actswithin physiologically-relevant concentrations motivated a search forother nitrated lipids that might serve related signaling actions.Herein, fatty acid nitroalkene products in plasma and urine are abundantand ubiquitous are reported. Of the total fatty acid content in redcells, linoleic acid and oleic acid comprise ˜8% and ˜18%, respectively(20). Thus, due to its prevalence and structural simplicity, oleic acidwas evaluated as a potential candidate for nitration. Herein, reportedis the synthesis, structural characterization and cell signalingactivity of 9- and 10-nitro-9-cis-octadecaenoic acids (nitrated oleicacid; OA-NO₂; FIG. 10) and that OA-NO₂ regioisomers are present in humanblood at levels exceeding those of LNO₂. Furthermore, OA-NO₂ activatesPPARγ with a greater potency than LNO₂. These data reveal that nitratedunsaturated fatty acids represent a novel class of lipid-derived,receptor-dependent signaling mediators.

Methods

Materials.

9-Octadecenoic acid (oleic acid) and its respective methyl ester,methyl-9-octadecenoate was purchased from Nu-Check Prep (Elysian,Minn.). LNO₂ and [¹³C]LNO₂ were synthesized as previously described(9,12); OA-NO₂ and [¹³C]OA-NO₂ were synthesized as described in below.Phenylselenium bromide, HgCl₂, NaNO₂, anhydrous tetrahydrofuran (THF),CH₃CN, CDCl₃, insulin, dexamethasone and 3-isobutyl-1-methylxanthinewere obtained from Sigma/Aldrich (St Louis, Mo.). Silica gel G and HFthin layer chromatography plates (250 and 2000 μm) were from Analtech(Newark, Del.). Methanolic BF₃, horseradish peroxidase-linked goatanti-rabbit IgG and Coomasie Blue were from Pierce (Rockford, Ill.).Synthetic solvents were of HPLC grade or better from Fisher Scientific(Fairlawn, N.J.). Solvents used for extractions and mass spectrometricanalyses were from Burdick and Jackson (Muskegon, Mich.). [¹³C]Oleicacid and [¹³C]linoleic acid were purchased from Cambridge IsotopeLaboratories, Inc. (Andover, Mass.). Anti-PPARγ and anti-β-actinantibodies were from Santa Cruz (Santa Cruz, Calif.); anti-aP2 antibodywas from Chemicon International Inc. (Temecula, Calif.).

Synthesis of OA-NO₂.

Oleic acid and [¹³C]oleic acid were nitrated as described (9,12), withmodifications. Briefly, oleic acid, HgCl₂, phenylselenium bromide andNaNO₂ (1:1.3:1:1, mol/mol) were combined in THF/acetonitrile (1:1, v/v)with a final concentration of 0.15 M oleic acid. The reaction mixturewas stirred (4 h, 25° C.), followed by centrifugation to sediment theprecipitate. The supernatant was recovered, the solvent evaporated invacuo, the product mixture redissolved in THF (original volume) and thetemperature reduced to 0° C. A ten-fold molar excess of H₂O₂ was slowlyadded with stirring to the mixture, which was allowed to react in an icebath for 20 min followed by a gradual warming to room temp (45 min). Theproduct mixture was extracted with hexane, the organic phase collected,the solvent removed in vacuo and lipid products solvated in CH₃OH.OA-NO₂ was isolated by preparative TLC using silica gel HF platesdeveloped twice in a solvent system consisting of hexane/ether/aceticacid (70:30:1, v/v). The region of silica containing OA-NO₂ was scrapedand extracted (21). Based on this synthetic rationale, two regioisomersare generated: 9- and 10-nitro-9-cis-octadecenoic acids (genericallytermed OA-NO₂). Thin layer chromatography does not resolve the twoisomers. [¹³C]OA-NO₂ was synthesized using [¹³C]oleic acid as areactant. Stock concentrations of OA-NO₂ isomers were quantitated bychemiluminescent nitrogen analysis (Antek Instruments, Houston, Tex.),using caffeine as a standard. All standards were diluted in methanol,aliquoted and stored under argon gas at −80° C. Under these conditions,OA-NO₂ isomers remain stable for >3 months.

OA-NO₂ Spectrophotometric Characterization.

OA-NO₂ stock solution concentrations were initially determined bychemiluminescent nitrogen analysis. These data were utilized todetermine dilution concentrations for subsequent spectral analysis. Anabsorbance spectrum of OA-NO₂ from 200-450 nm was generated using 23 μMOA-NO₂ in phosphate buffer (100 mM, pH 7.4) containing 100 μM DTPA. Theextinction coefficients (γ) for OA-NO₂ and the isotopic derivative[¹³C]OA-NO₂ were measured (λ₂₇₀) using a UV-VIS spectrophotometer(Shimadzu, Japan). Absorbance values for increasing concentrations ofOA-NO₂ and [¹³C]OA-NO₂ were plotted against concentration to calculateε.

NMR Spectrometric Analysis of OA-NO₂

¹H and ¹³C NMR spectra were measured using a Varian INOVA 300 and 500MHz NMR and recorded in CDCl₃. Chemical shifts are in δ units (ppm) andreferenced to residual proton (7.26 ppm) or carbon (77.28 ppm) signalsin deuterated chloroform. Coupling constants (J) are reported in Hertz(Hz).

Gas Chromatography Mass Spectrometric Characterization of OA-NO₂.

Methyl esters of purified synthetic OA-NO₂ regioisomers were analyzed byelectron impact ionization gas chromatography mass spectrometry (EIGC-MS). Fatty acid methyl esters of OA-NO₂ were synthesized, extractedwith hexane, washed twice with saturated saline, redissolved in undecaneand analyzed by GC-MS using a Saturn 2000 Tandem Mass Spectrometercoupled to a Varian 3800 Gas Chromatograph. Samples were ionized byelectron impact at +70 eV. The methyl ester-derivatized OA-NO₂regioisomers were resolved using a 30 m capillary column (5% phenyl 95%dimethylpolysiloxane; CP-Sil 8CB-MS, Varian) with the followingtemperature gradient: 80° C. (2 min); 80 to 170° C. at 20° C./min; 170to 240° C. at 2° C./min and 240 to 280° C. at 5° C./min. Helium was usedas a carrier gas.

Structural Characterization of OA-NO₂ by Electrospray Ionization TripleQuadrupole Mass Spectrometry (ESI MS/MS).

Qualitative analysis of OA-NO₂ by ESI MS/MS was performed using a hybridtriple quadrupole-linear ion trap mass spectrometer (4000 Q trap,Applied Biosystems/MDS Sciex). To characterize synthetic and endogenousOA-NO₂, a reverse-phase HPLC separation was developed using a 150×2 mmC18 Phenomenex Luna column (3 μm particle size). Lipids were eluted fromthe column using a gradient solvent system consisting of A (H₂Ocontaining 0.1% NH₄OH) and B (CNCH₃ containing 0.1% NH₄OH) under thefollowing conditions: 20 to 65% B (10 min); 65 to 95% B (1 min; hold for3 min) and 95 to 20% B (1 min; hold for 3 min). Using these gradientconditions, OA-NO₂ elutes after LNO₂ positional isomers. OA-NO₂ wasdetected using a multiple reaction monitoring (MRM) scan mode byreporting molecules that undergo a m/z 326/279 mass transition, which isconsistent with the loss of the nitro group ([M-(HNO₂)]⁻). Concurrentwith MRM determination, enhanced product ion analysis (EPI) wasperformed to generate characteristic and identifying fragmentationpatterns of eluting species with a precursor mass of m/z 326. EPIanalysis utilizes the trap functionality of the triple quadrupole to“concentrate” fragment ions to enhance sensitivity. Zero grade air wasused as source gas, and nitrogen was used in the collision chamber.

Red Blood Cell Isolation and Lipid Extraction.

Peripheral blood from fasting healthy human volunteers was collected byvenipuncture into heparinized tubes (UAB Institutional ReviewBoard-approved protocol #X040311001). Blood was centrifuged (1200×g; 10min), the buffy coat removed and erythrocytes were isolated. Lipidextracts were prepared from red cells and plasma (21) and directlyanalyzed by mass spectrometry. Care was taken to avoid acidificationduring extraction to prevent artifactual lipid nitration due to thepresence of endogenous nitrite (9). In experiments using urine as thebiological specimen (UAB Institutional Review Board-approved protocol#X040311003), extraction conditions were identical.

Detection and Quantitation of OA-NO₂ in Human Blood and Urine.

Quantitation of OA-NO₂ in biological samples was performed as described(9), with modifications. Matched blood and urine samples were obtainedafter >8 hr fasting; urine was collected from the first void of the day.During the monophase stage of the lipid extraction (21), [¹³C]OA-NO₂ wasadded as internal standard to correct for losses due to extraction.Nitrated fatty acids were then analyzed by HPLC ESI MS/MS. Lipids wereeluted from the HPLC column using an isocratic solvent system consistingof CH₃CN:H₂O:NH₄OH (85:15:0.1, v/v), resulting in the co-elution of thetwo OA-NO₂ regioisomers. During quantitative analyses, two MRMtransitions were monitored: m/z 326/279 (OA-NO₂) and m/z 344/297([¹³C]OA-NO₂), transitions consistent with the loss of the nitro groupfrom the respective precursor ions. The areas under each peak wereintegrated, the ratio of analyte to internal standard areas wasdetermined and levels of OA-NO₂ were quantitated using Analyst 1.4quantitation software (Applied Biosystems/MDS Sciex) by fitting the datato an internal standard curve. Data are expressed as mean±std dev (n=10;5 female and 5 male).

Qualitative Analysis of Nitro- and Nitrohydroxy-Adducts of Fatty Acids.

Using HPLC ESI-MS/MS, blood and urine samples were evaluated for thepresence of allylic nitro derivatives other than LNO₂ and OA-NO₂. HPLCseparations were performed similarly to those used to characterizeOA-NO₂, with some modifications. Alternative MRM transitions were usedto detect other potential nitroalkene derivatives. Based on what appearsto be a common fragmentation product of nitrated fatty acids (i.e., lossof the nitro group; [M−HNO₂]⁻), theoretical MRM transitions weredetermined for nitrated linolenic (18:3-NO₂), arachidonic (20:4-NO₂) anddocosahexaenoic acids (22:6-NO₂). MRM transitions for nitrohydroxyadducts were also monitored: 18:1(OH)—NO₂; 18:2(OH)—NO₂; 18:3(OH)—NO₂;20:4(OH)—NO₂ and 22:6(OH)—NO₂.

LNO₂ and OA-NO₂ Decay.

The relative rates of LNO₂ and OA-NO₂ decay in aqueous solution weredetermined by incubating 3 μM LNO₂ and OA-NO₂ in phosphate buffer (100mM, pH 7.4, 37° C.). During the two hour incubation, aliquots wereremoved and analyzed for LNO₂ and OA-NO₂ content. The aliquots wereextracted as described (21) and 1 μM [¹³C]LNO₂ was added during themonophase stage of the extraction procedure as an internal standard.Non-decayed LNO₂ and OA-NO₂ were quantitated via HPLC ESI-MS/MS asdescribed above.

PPAR Transient Transfection Assay.

CV-1 cells from the ATCC (Manassas, Va.) were grown to ˜85% confluencein DMEM/F12 supplemented with 10% FBS, 1% penicillin-streptomycin.Twelve hours before transfection, the media was removed andantibiotic-free media was applied. Cells were transiently co-transfectedwith a plasmid containing the luciferase gene under the control of threetandem PPAR response elements (PPRE) (PPRE×3 TK-Luciferase) and PPARγ,PPARα or PPARδ expression plasmids, respectively. In all cases,fluorescence protein (GFP) expression plasmid was co-transfected as thecontrol for transfection efficiency. Twenty-four hours aftertransfection, cells the returned to OptiMEM (Invitrogen, Carlsbad,Calif.) for 24 hr and then treated as indicated for another 24 hr.Reporter luciferase assay kits from Promega (Madison, Wis.) were used tomeasure the luciferase activity according to the manufacturer'sinstructions with a luminometer (Victor II, Perkin-Elmer). Luciferaseactivity was normalized by GFP units. Each condition was performed intriplicate in each experiment (n>3).

3T3-L1 Differentiation and Oil Red O Staining.

3T3-L1 preadipocytes were propagated and maintained in DMEM containing10% FBS. To induce differentiation, 2-day post-confluent preadipocytes(designated day 0) were cultured in DMEM containing 10% FBS plus 1 and 3μM OA-NO₂ for 14 days. The medium was changed every two days.Rosiglitazone (3 μM) and oleic acid (3 μM) were used as positive andnegative controls, respectively. Differentiated adipocytes were stainedwith oil red O as previously (22).

[³H]-2-Deoxy-D-Glucose Uptake Assay in Differentiated 3T3-L1 Adipocytes.

[³H]-2-deoxy-D-glucose uptake was analyzed as previously (23). 3T3-L1preadipocytes were grown in 24-well tissue culture plates, 2-daypost-confluent monolayers were treated with 10 μg/ml insulin, 1 μMdexamethasone, and 0.5 mM 3-isobutyl-1-methylxanthine in DMEM containing10% FBS for two days, then cells were maintained in 10 μg/ml insulin inDMEM containing 10% FBS for 6 days (medium was changed every threedays). Eight days after induction of adipogenesis, test compounds inDMEM containing 10% FBS were added for an additional 2 days (medium waschanged every day). The PPARγ-specific antagonist GW9662 was added 1 hrbefore other additions. After two rinses with serum-free DMEM, cellswere incubated for 3 hr in serum-free DMEM and rinsed at roomtemperature three times with freshly prepared KRPH buffer (5 mMphosphate buffer, 20 mM HEPES, 1 mM MgSO₄, 1 mM CaCl₂, 136 mM NaCl, 4.7mM KCl, pH 7.4). The buffer was replaced with 1 μCi/ml of[³H]-2-deoxy-D-glucose in KRPH buffer for 10 min at room temperature.Cells were then rinsed three times with cold PBS, lysed overnight in 0.8N NaOH (0.4 ml/well), neutralized with 26.6 μl of 12 N HCl and 360 μl oflysate was added into 4 ml Scinti-safe plus TM 50% for radioactivitydetermination by liquid scintillation counting.

Results

Detection and Identification of Nitrated PUFA.

The discovery that LNO₂ is present in vivo motivated a search foradditional endogenous nitrated fatty acids that may also act as lipidsignaling molecules. To survey plasma and urine for other nitrated fattyacids, lipid extracts from healthy human blood donors were analyzed byHPLC ESI MS/MS in the multiple reaction monitoring (MRM) scan mode. MRMtransitions were calculated for the nitro- and nitrohydroxy-adducts ofsix fatty acids, as shown in Table 2, and were used to detect nitro- andnitrohydroxy-adducts present in plasma and urine lipid extracts using anisocratic HPLC elution methodology (FIG. 11). Gradient elution methodsusing MRM and EPI scan modes were used for structural confirmation (notshown). Due to the lack of appropriate stable isotope internal standardsfor all derivatives, data are presented as base-peak spectra and arequalitative evidence that these species exist in vivo. Massspectrometric analysis revealed that nitrated adducts of all monitoredunsaturated fatty acids are present in blood and urine, which includethe Michael-like addition products of these species with H₂O detected asnitrohydroxy adducts. Owing to its predominant abundance and structuralsimplicity, oleic acid was synthesized as a standard to specificallyquantitate endogenous OA-NO₂ content and signaling activity.

TABLE 2 Multiple reaction monitoring (MRM) transitions for fatty acidnitroalkene derivatives MRM values for nitroalkene andnitrohydroxy-adducts of fatty acids were based on the common loss of thenitro group that occurs during collision-induced dissociation ofnitrated fatty acids. Carbons: Nitro Nitrohydroxy double adduct adductFatty Acid bonds (—NO₂) (L(OH)—NO₂) Oleic 18:1 326/279 344/297 Linoleic18:2 324/277 342/295 Linolenic 18:3 322/275 340/293 Arachidonic 20:4348/301 366/319 Eicosapentaenoic 20:5 346/299 364/317 Docosahexaenoic22:6 372/325 390/343

Synthesis and Purification of OA-NO₂.

Nitration of oleic acid by nitrosenylation will yield two potentialregioisomers of OA-NO₂ (FIG. 10). Upon purification, analytical TLC, GCand LC-mass spectrometry of synthetic OA-NO₂ indicated no contaminationby oleic acid nor oxidized products (not shown).

NMR Analysis of OA-NO₂.

The structure of synthetic OA-NO₂ (a 1:1 mixture of C9- and C10regioisomers) was analyzed by ¹H and ¹³C NMR. NMR splitting patterns aredesignated as s, singlet; d, doublet; t, triplet; q, quartet; m,multiplet and br, broad. ¹H-NMR (CDCl₃): δ 11.1 (br s, 1H); 7.06 (dd,1H, J=7.8 Hz); 3.75 (t, 2H, J=6.7 Hz); 2.55 (t, 2H, J=7.6 Hz); 2.36 (q,2H, J=7.6 Hz); 2.33 (m, 2H); 2.20 (q, 2H, J=7.3 Hz); 1.85 (m, 2H); 1.61(m, 4H); 1.47 (m, 4H); 1.32-1.25 (m, 8H); 0.87 (dt, 3H, J=7.0 Hz). The¹H spectrum and proposed assignments of diagnostic peaks are presentedin FIG. 12A: 11.1 (COOH); 7.06 (C9 or C10, alkene proton, each a tripletfrom coupling to neighboring methylene —CH₂, with regioisomerssuperimposed on each other, appearing on one NMR spectrometer as adoublet of triplets and on the other as a quartet, which is really asuperimposed pair of triplets); 3.75 (C8 or C11, allylic methyleneneighboring nitro group; nitroalkene more electron-withdrawing thancarbonyl); 2.55 (C2 methylene neighboring carbonyl); 2.20 (C8 or C11,allylic methylene); 1.85 (C7 or C12, methylene next to nitro-allylposition); 1.61 (C3 methylene); 0.87 (C18 terminal methyl, superimposedregioisomers resulting in doublet of triplets). There is no indicationof oleic acid or synthetic intermediates and no trans-isomers of OA-NO₂in the proton spectrum. The local environment of each proton in the 9-and 10-nitrated cis-isomers is not distinguishable, so each signal isindicative of relative positions in the alkene region, and absolutepositions in the extreme portions of the molecule (i.e., the terminalmethyl and acid groups). The aliphatic regions are too similar for exactassignments.

Further structural characterization was performed by ¹³C NMR. From thespectrum, 30 total peaks were observed: δ180.1, 180.0; 152.2, 152.0;136.8, 136.4; 68.1; 34.2; 32.0; 29.6; 29.5; 29.5; 29.5; 29.4; 29.4;29.3; 29.2; 29.1; 28.8; 28.7; 28.3; 28.2; 28.1; 28.1; 26.6; 26.6; 25.8;24.8; 22.9; 14.3. The proposed assignment of diagnostic peaks ispresented in FIG. 12B: 180.1, 180.0 (C1 carbonyl); 152.2, 152.0 (C9 orC10, alkene attached to NO₂); 136.7, 136.4 (C9 or C10, alkene); 68.2 (C8or C11, CH₂ allylic to NO₂); 34.2 (C2, CH₂ neighboring carbonyl); 32.1(C8 or C11, allylic distal to NO₂); 14.3 (C18, terminal CH₃). The totalintegrated signal is less than the total number of carbons for the twoisomers (i.e., 30 vs. 36); thus, some peaks are identifiable doubletsfrom isomers (e.g., carbonyl peaks), whereas others areindistinguishable and appear as a singlet (e.g., the methyl peak).

Spectral Characterization of OA-NO₂.

The spectrum of OA-NO₂ was acquired in phosphate buffer in the presenceof the iron chelator DTPA (FIG. 13A). An absorbance maximum at 270 nmwas identified that is ascribed to electron absorption by the NO₂ group.Extinction coefficients for OA-NO₂ and [¹³C]OA-NO₂ were determined byplotting absorbance (λ₂₇₀) vs. concentration, giving m=AU·cm⁻¹·mM⁻¹ anda calculated ε=8.22 and 8.23 cm⁻¹ mM⁻¹, for OA-NO₂ and [¹³C]OA-NO₂,respectively (FIG. 13B).

Characterization of OA-NO₂ by GC-MS/MS.

Capillary columns used for gas chromatography have high resolvingcapacity and can separate regio- and diastereoisomers. Thus, initialmass spectrometric characterization of OA-NO₂ regioisomer methyl esterswas performed by GC-MS (FIG. 14). Filtering total ion count (TIC) datato show ions with m/z 342 ([M+H]⁺) and 306 (source fragment of m/z 342;[M-2H₂O]⁺) reveals two peaks appearing at 37.4-37.7 min (FIG. 14A, upperpanel). Product ion analysis of these peaks generated an identifyingspectrum for each species (FIG. 14B). Ions with m/z 324 and 306represent the loss of one and two H₂O from the parent compound. Theunique product ions m/z 168 and 156 for the first and second elutingpeaks, respectively, were chosen as identifiers of each regioisomer.Filtering the MS/MS TIC for these product ions generated chromatogramsthat show the individual regioisomers of OA-NO₂ (FIG. 14A, middle andlower panels).

Characterization and Quantitation of OA-NO₂ by ESI-MS/MS.

Using the gradient HPLC protocol described in Methods, synthetic OA-NO₂regioisomers eluted from the reverse phase column as two overlappingpeaks (FIG. 15). The HPLC elution profiles for synthetic OA-NO₂ and[¹³C]OA-NO₂ were virtually identical (FIG. 15A, left panels). Concurrentproduct ion analysis of the overlapping peaks showed spectra consistentwith OA-NO₂-derived species (FIG. 15A, right panels), with majorfragments identified in Table 3. Using these same parameters and machinesettings, lipid extracts of packed red cells and plasma were analyzed(FIG. 15B). The product ion spectra for OA-NO₂ of red cells and plasmaare identical to those obtained from synthetic OA-NO₂, revealing thatOA-NO₂ is endogenously present in blood. Interestingly, the HPLC elutionprofiles for plasma- and blood-derived OA-NO₂ show single peaks ratherthan overlapping species, suggesting only one regio-isomer is present invivo.

TABLE 3 Collision-induced dissociation fragments of nitroalkene fattyacid derivatives Nitroalkene derivatives of fatty acids were analyzed byelectrospray-ionization tandem mass spectrometry. Product ion spectrafrom synthetic standards were obtained in the negative ion mode asdescribed in Experimental Procedures. Major fragments generated for eachstandard are listed below. Mass/charge (m/z) OA-NO₂ [¹³C]OA-NO₂ LNO₂ 344— [M − H] — 326 [M − H] [M − H₂O] — 324 — — [M − H₂O] 308 [M − H₂O] [M −2H₂O] — 306 — — [M − H₂O] 295 — [M − HNO₂] — 293 — — [M − HNO] 279 [M −HNO₂] — — 277 — — [M − HNO₂] 264 — [M − (2H₂O + CO₂)] — 246 [M − (2H₂O +CO₂)] — — 244 — — [M − (2H₂O + CO₂)]

To quantitate OA-NO₂ content in red cells and plasma, lipid extractswere separated using an isocratic HPLC elution protocol wherein analytesco-elute at 2 min; MRM transitions for OA-NO₂ and [¹³C]OA-NO₂ weremonitored (not shown). The concentration of OA-NO₂ in biological sampleswas determined from the ratio of analyte to internal standard peak areasusing an internal standard curve that is linear over four orders ofmagnitude. The limit of quantitation (LOQ; determined as ten times thestandard deviation of the noise) was calculated to be ˜1.2 fmol oncolumn (not shown). Blood samples obtained from ten healthy humanvolunteers (5 female, 5 male, ages ranging from 24 to 51) revealed freeOA-NO₂ in red cells (i.e., OA-NO₂ not esterified to glycerophospholipidsor neutral lipids) to be 59±11 pmol/ml packed cells (Table 4). Totalfree and esterified OA-NO₂, the amount present in saponified samples,was 214±76 pmol/ml packed cells. Thus, ˜75% of OA-NO₂ in red cellsexists as esterified fatty acids (9). In plasma, the free and esterifiedOA-NO₂ concentrations were 619±52 and 302±369 nM, respectively, and wereobserved to be more abundant than linoleic acid nitration products (9).

TABLE 4 Nitrated Oleic Acid in human blood a comparison with nitratedlinoleic acid Venous blood was obtained from healthy human volunteersand centrifuged, and plasma and red cells were extracted and preparedfor mass spectrometric analysis as described in Experimental Procedures.During sample preparation, [¹³C]OA-NO₂ was added as an internal standardto correct for losses. OA-NO₂ was quantitated by fitting analyte tointernal standard area ratios obtained by MS to an internal standardcurve. Concentration values for LNO₂ in the vascular compartment wereobtained from ref. 9. Data are expressed as mean ± std dev (n-10; 5female and 5 male) Compartment Fraction [OA-NO₂] (nM) [LNO₂] (nM) PlasmaFree 619 ± 52  79 ± 35 Esterified 302 ± 369 550 ± 275 Total 921 ± 421630 ± 240 Packed red cells Free 59 ± 11 50 ± 17 Esterified 155 ± 65  199± 121 Total 214 ± 76  249 ± 104 Whole Blood* Total  639 ± 366*  477 ±128* *Assuming a 40% hematocrit

Characterization of Nitrohydroxy Allylic Derivatives.

The initial survey to detect nitrated fatty acids indicated thatnitrohydroxy allylic derivatives are also present in plasma and urine(FIG. 11). Their presence was confirmed by product ion analysis runconcomitantly with MRM detection (FIG. 16). Structures of possibleadducts are presented with diagnostic fragments and product ion spectrafor 18:1(OH)—NO₂, 18:2(OH)—NO₂ and 18:3(OH)—NO₂. Both the 9- and10-nitro regioisomers of 18:1(OH)—NO₂ are present in urine (A) andplasma (not shown), as evidenced by the intense peaks corresponding tom/z 171 and 202 (A). Also present are fragments consistent with the lossof a nitro group and water (m/z 297 and 326, respectively). The production spectrum obtained from 18:2(OH)—NO₂ shows a predominant fragment(m/z 171), consistent with an oxidation product of LNO₂ nitrated at the10-carbon (B). Diagnostic fragments for the three other potentialregioisomers were not apparent. Finally, multiple regioisomers of18:3(OH)—NO₂ are present (C). For all three Δ-9 fatty acids monitored,the predominant fragment generated during collision-induced dissociation(CID) is m/z 171.

Activation of PPARs by OA-NO₂.

Recently, LNO₂ has been identified as an endogenous PPAR ligand (17).Considering the high levels of OA-NO₂ detected in vivo, OA-NO₂ wasevaluated and compared with LNO₂ as a potential ligand for PPARα, PPARγand PPARδ. CV-1 cells were transiently co-transfected with a plasmidcontaining the luciferase gene under regulation by three PPAR responseelements (PPRE) in concert with PPARγ, PPARα or PPARδ expressionplasmids. Dose-dependent activation by OA-NO₂ was observed for allPPARs, (FIG. 17A), with PPARγ showing the greatest response (significantactivation at 100 nM); PPARα and PPARδ showed significant activation at˜300 nM OA-NO₂. Nitrated oleic acid was consistently more potent thanLNO₂ in the activation of PPARγ, with 1 μM OA-NO₂ typically inducing thesame degree of reporter gene expression as 3 μM LNO₂ or 1 μMRosiglitazone; these increases were partially inhibited by the PPARγantagonist GW9662 (FIG. 17B). Native fatty acids did not activate PPARsat these concentrations (not shown). The greater potency of OA-NO₂ as aPPARγ agonist compared to LNO₂ motivated evaluation of the relativestability of these molecules, because LNO₂ decays in aqueous milieu togenerate products that do not activate PPARs (17,24). Compared withLNO₂, OA-NO₂ is relatively stable, with only minimal decay occurringafter 2 hr; a time when ˜80% LNO₂ decay was noted (FIG. 17C).

The signaling actions of OA-NO₂ as a PPARγ ligand was assessed byevaluating its impact on adipocyte differentiation, as PPARγ-dependentgene expression plays an essential role in the development of adiposetissue (25,26). 3T3-L1 preadipocytes were treated with OA-NO₂ (1 μM),LNO₂ (3 μM) and negative controls for two weeks (FIG. 18A). Previously,LNO₂ was shown to induce pre-adipocyte differentiation to the sameextent as Rosiglitazone (17). Adipocyte differentiation was assessedboth morphologically and via oil red O staining, which revealed theaccumulation of intracellular lipids. Vehicle, oleic acid and linoleicacid did not induce adipogenesis. In contrast, OA-NO₂ and LNO₂induced >30% of 3T3-L1 preadipocyte differentiation. Rosiglitazone, asynthetic PPARγ ligand, also induced PPARγ-dependent preadipocytedifferentiation. OA-NO₂— and Rosiglitazone-induced pre-adipocytedifferentiation resulted in expression of specific adipocyte markers(PPARγ2 and aP2); oleic acid had no effect on these gene products (FIG.18B). PPARγ ligands also play a central role in glucose uptake andmetabolism, with agonists widely used as insulin-sensitizing drugs.Consistent with its potent PPARγ ligand activity, OA-NO₂ induced anincrease in differentiated adipocyte glucose uptake (FIG. 19A). Thiseffect of OA-NO₂ was paralleled by higher concentrations of LNO₂ (3 μM).The increased adipocyte glucose uptake, induced by nitrated fatty acidsand the positive control Rosiglitazone, was partially inhibited byGW9662 (FIG. 19B). In aggregate, these observations reveal that OA-NO₂manifests well-characterized PPARγ-dependent signaling actions.

Discussion

The nitration of hydrocarbons has long been recognized (27). Followingthe more recent discovery of vascular cell signaling actions of oxidesof nitrogen (1,28), it is now also appreciated that .NO-derived speciesmediate oxidation, nitrosation, nitrosylation and nitration reactions ofprotein, DNA and unsaturated fatty acids (29). These reactionsfrequently yield stable products that influence target moleculestructure and function to either a) translate the signaling actions of.NO or b) mediate pathogenic responses when occurring in “excess”.

The reactions of .NO and its redox-derived products with lipids ismultifaceted. Model studies of photochemical air pollutant-induced lipidoxidation revealed that exceedingly high concentrations of nitrogendioxide (.NO₂) could induce nitration of fatty acids inphosphatidylcholine liposomes and fatty acid methyl ester preparations(30-32). Subsequently, reaction systems designed to model theinteractions of endogenous .NO and .NO-derived species [e.g.,peroxynitrite (ONOO⁻) and nitrous acid (HNO₂)] with fatty acids showedthat a) .NO mediates potent inhibition of autocatalytic radical chainpropagation reactions of lipid peroxidation (33,34) and b) .NO-derivedspecies produce both nitrated and oxidized derivatives of unsaturatedfatty acids (3,35). One product of these reaction pathways, LNO₂, ispresent at ˜500 nM concentration in healthy human red cells and plasmaand serves as a ligand for the PPAR nuclear lipid receptor family(9,17). This insight, coupled with the fact that oleic acid is the mostabundant unsaturated fatty acid in mammals and plants, motivated thepresent search for other potential endogenous nitrated fatty acidderivatives that might translate tissue redox signaling reactions.

The structure of OA-NO₂ (FIG. 10) was defined on the basis of thesynthetic rationale and NMR analysis (FIG. 12). Proton and ¹³C NMRspectra indicate that the synthetic OA-NO₂ is comprised of tworegioisomers, 9- and 10-nitro-9-cis-octadecenoic acids, with notrans-isomers apparent. Peaks characteristic of the nitro-allylic andallylic carbons in the ¹³C spectrum both appear as doublets that areequal in intensity, indicating an equivalent distribution betweenregioisomers. EI GC MS confirmed the presence of 9- and10-nitro-9-cis-octadecenoic acids by mass and differential retentiontimes (FIG. 14). HPLC ESI MS/MS was used to further characterizesynthetic OA-NO₂. The combined fragmentation pattern of OA-NO₂regioisomers was obtained by CID, which provided a “molecularfingerprint” that was used to identify OA-NO₂ in biological samples(FIG. 15). ESI MS/MS analysis of lipid extracts derived from plasma andred cells yielded spectra with identical HPLC retention times and majorproduct ions, confirming that OA-NO₂ exists endogenously; however, it isnot possible from MS analysis to determine the cis/trans conformation ofregioisomers. Quantitative analysis of plasma and red cells revealedthat OA-NO₂ is present in the vasculature at net concentrations ˜50%greater than LNO₂ (Table 4). Combined, esterified and free OA-NO₂ andLNO₂ are well above 1 μM, a concentration range capable of elicitingcell signaling responses.

The nitro functional groups of OA-NO₂ and LNO₂ are located on olefiniccarbons. This configuration imparts a unique chemical reactivity thatenables the release of .NO during aqueous decay of these nitroalkenederivatives via a modified Nef reaction (24). Furthermore, the α-carbonproximal to the alkenyl nitro group is strongly electrophilic, whichreadily reacts with H₂O via a Michael addition mechanism to generatenitrohydroxy adducts (FIGS. 11 and 16). Nitrohydroxy-arachidonic acidspecies have previously been detected in bovine cardiac muscle (36), andnitrohydroxy-linoleic acid has been identified in lipid extractsobtained from hypercholesterolemic and post-prandial human plasma,suggesting that this is a ubiquitous derivative (37). The presentidentification of a wide spectrum of nitrated fatty acids andcorresponding nitrohydroxy-fatty acid derivatives in human plasma andurine reveals that nitration reactions occur with all unsaturated fattyacids (FIGS. 11, and 16). The hydroxyl moiety of nitrohydroxy-fattyacids destabilizes the adjacent carbon-carbon bond, resulting inheterolytic scission reactions that a) generate predictable fragmentsduring CID (FIG. 16) and b) render potentially unique cell signalingactions to fatty acid nitrohydroxy derivatives. Present data indicates,however, that nitrohydroxy adducts of LNO₂ and OA-NO₂ are not avidligands for PPARγ (FIG. 17C).

Multiple mechanisms can support the basal and inflammatory nitration offatty acids by .NO-derived species, including .NO₂-initiatedauto-oxidation of polyunsaturated fatty acids via hydrogen abstractionfrom the bis-allylic carbon and nitration by acidified NO₂ ⁻ (31,38-42). Of relevance to both basal and inflammatory cell signaling, .NO₂can be derived from multiple reactions. This includes the homolyticscission of both peroxynitrous acid (ONOOH) and nitrosoperoxocarbonate(ONOOCO₂ ⁻), as well as the oxidation of NO₂ ⁻ by heme peroxidases(43,44). Present data supports that all of these alkenyl nitrationmechanisms can yield nitrated fatty acids that are structurally similaror identical to the OA-NO₂ and LNO₂ detected clinically. Nitration by afree radical mechanism might suggest that all olefinic carbons within afatty acid would be susceptible nitration targets; with the additionallikelihood of double bond rearrangement and conjugation. The discoveryof OA-NO₂ lends critical perspective to this issue, becausemonounsaturated fatty acids are less susceptible to freeradical-mediated hydrogen abstraction reactions. In view of the datapresented herein, alternative fatty acid nitration mechanisms may thusalso be viable. For example, nitration by an ionic addition reaction(e.g., nitronium ion, NO₂ ⁺) can generate singly nitrated fatty acidswith no double bond-rearrangement. Since NO₂ ⁺ readily reacts with H₂O,this species may require stabilization in the hydrophobic milieu of themembrane bilayer or localized catalysis (e.g., reaction of ONOO— withtransition metals) to serve as a biologically-relevant nitratingspecies. Finally, radical addition reactions may also occur with mono-and polyunsaturated fatty acids to yield non-conjugated nitroalkenederivatives of polyunsaturated fatty acids, as revealed by studies ofacidified NO₂ ⁻ and .NO₂-mediated fatty acid methyl ester oxidation andnitration profiles (32,41).

Of important relevance to mechanisms underlying fatty acid nitration invivo, the nitrohydroxy adducts of Δ9 unsaturated fatty acids examined inthe present study (18:1, 18:2 and 18:3) all have a predominant fragmentof m/z 171 (FIG. 16). This mass is consistent with 9-oxo-nonanoic acid,a fragment generated with authentic standards when the nitro group islocated at the 10-carbon and the hydroxy moiety at the 9-carbon (datanot shown). This suggests either strict steric control or enzymaticmechanisms regulating the stereospecificity of biological fatty acidnitration. The nitration of Δ9 unsaturated fatty acids to C10nitroalkene derivatives, with retention of double bond arrangement,supports that stereospecific enzymatic reactions may mediate fatty acidnitration. It is also possible that nitrated fatty acids are madebioavailable from dietary sources that give rise to specific fatty acidnitroalkene derivatives.

Designation of nitroalkene derivatives as a class of signaling moleculesis contingent upon ascribing specific bioactivities to multiple memberswithin the class at clinically-relevant concentrations. Nitrolinoleatehas been observed to inhibit neutrophil and platelet function viacGMP-independent, cAMP-mediated mechanisms (10-12). Also, aqueous decayof LNO₂ yields .NO, a reaction that is facilitated by translocation ofLNO₂ from a hydrophilic to hydrophobic microenvironment, which in turninduces cGMP-dependent vessel relaxation (12,24). Recently, LNO₂ hasalso been reported to serve as a robust ligand for PPARγ (17), a nuclearhormone receptor that binds lipophilic ligands to induce DNA binding ofthe transcription factor complex at DR1-type motifs in the promotersites of target genes. Downstream effects of PPARγ activation includemodulation of metabolic and cellular differentiation genes andregulation of inflammatory responses, adipogenesis and glucosehomeostasis (45,46). In the vasculature, PPARγ is expressed inmonocytes, macrophages, smooth muscle cells and endothelium (47) andplays a central role in regulating the expression of genes related tolipid trafficking, cell proliferation and inflammatory signaling. Hereinwe show that OA-NO₂ also serves as a PPARγ, α and δ ligand that rivalsor exceeds the potency of LNO₂ and synthetic PPAR ligands such asfibrates and thiazolidinediones (FIGS. 17-19). The increased potency ofOA-NO₂ as a PPARγ ligand is either due to increased aqueous stabilityrelative to LNO₂ (FIG. 17C) or increased receptor affinity. The combinedblood concentrations of OA-NO₂ and LNO₂ in healthy humans exceeds 1 μM(Table 4), concentrations well-within the range needed to activate PPARreceptors and exceeding those of previously-proposed endogenous PPARγligands. These observations have broad implications for the .NO andredox signaling reactions that play a crucial role in dysregulated cellgrowth and differentiation, metabolic syndrome, atherosclerosis anddiabetes—all clinical pathologies that also include a significantcontribution from PPAR-regulated cell signaling mechanisms (48).

The regulation of inflammation by inhibiting eicosanoid synthesis is awell-established and prevalent target of anti-inflammatory drugstrategies. Much less well-understood are the concerted cell signalingmechanisms by which inflammation is favorably resolved in vivo. Whilethe integrated in vivo tissue signaling activities of nitrated fattyacids remain to be defined, studies to date indicate that thesepluripotent signaling mediators generally manifest salutary metabolicand anti-inflammatory actions (10-12,17). The capability ofredox-derived lipid signaling molecules to mediate the resolution ofinflammation is a relatively new concept, with lipoxins representing onenew class of lipid mediators that may also act in this manner (49).Endogenous concentrations of OA-NO₂ and LNO₂ are abundant and areincreased by oxidative inflammatory reactions. Thus, nitrated fattyacids are expected to play both receptor-dependent (via PPAR ligandactivity) and cyclic nucleotide-mediated roles in transducing the redoxsignaling actions of oxygen and .NO, thereby regulating organ function,cell differentiation, cell metabolism and systemic inflammatoryresponses.

Throughout Example 3, several publications have been referenced. Thesepublications are listed in the Reference List for Example 3. Thedisclosures of these publications in their entireties are herebyincorporated by reference into this application in order to more fullydescribe the compounds, compositions and methods described herein.

REFERENCE LIST FOR EXAMPLE 3 Reference List

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Example 4

The aqueous decay and concomitant release of nitric oxide (.NO) bynitrolinoleic acid (10-nitro-9,12-octadecadienoic acid and12-nitro-9,12-octadecadienoic acid; LNO₂) is reported. Massspectrometric analysis of reaction products support the Nef reaction asthe mechanism accounting for the generation of .NO by the aqueousreactions of fatty acid nitroalkene derivatives. Nitrolinoleic acid isstabilized by aprotic milieu, with LNO₂ decay and .NO release stronglyinhibited by phosphatidylcholine-cholesterol liposome membranes anddetergents when present at levels above their critical micellarconcentrations. The release of .NO from LNO₂ was induced by the UVphotolysis and I₃-based ozone chemiluminescence reactions currentlybeing used to quantify putative protein nitrosothiol (RSNO) andN-nitrosamine derivatives. This reactivity of LNO₂ complicates thequalitative and quantitative analysis of biological oxides of nitrogenwhen applying UV photolysis and triiodide (I₃ ⁻)-based analyticalsystems in biological preparations typically abundant in nitrated fattyacids. These results reveal that nitroalkene derivatives of linoleicacid are pluripotent signaling mediators that act via not onlyreceptor-dependent mechanisms, but also by transducing the signalingactions of .NO via pathways subject to regulation by the relativedistribution of LNO₂ to hydrophobic versus aqueous microenvironments.

Nitrolinoleic acid (10-nitro-9,12-octadecadienoic acid and12-nitro-9,12-octadecadienoic acid; abbreviated as LNO₂) is present inplasma lipoproteins and red blood cell membranes at concentrations of˜500 nM, rendering this species the most quantitatively abundantbiologically-active oxide of nitrogen in the human vascular compartment(1). Nitrolinoleic acid is a product of nitric oxide (.NO)-dependentlinoleic acid nitration reactions that predominantly occur at the C10and C₁₂ alkene carbons. The positional isomer distribution of the LNO₂alkenyl nitro group indicates that in vivo fatty acid nitration is aconsequence of nucleophilic (nitronium group, NO₂ ⁺) and/or radical(nitrogen dioxide, .NO₂) addition reactions with olefinic carbons.

Recent observations reveal that LNO₂ is a pluripotent signaling mediatorthat acts via both receptor-dependent and -independent pathways.Nitrated fatty acids are specific and high affinity endogenous ligandsfor peroxisome proliferator-activated receptors (2), and serve toactivate receptor-dependent gene expression at physiologicalconcentrations. LNO₂ also activates cAMP-dependent protein kinasesignaling pathways in neutrophils and platelets, serving todown-regulate the activation of these inflammatory cells (3,4). Finally,LNO₂ induces vessel relaxation in an endothelial-independent manner (5).This LNO₂-mediated relaxation of phenylephrine-preconstricted aorticrings was 1) a consequence of LNO₂-induced stimulation of smooth musclecell and aortic segment cGMP content, 2) inhibitable by the .NOscavenger oxyhemoglobin and 3) ODQ-inhibitable (e.g., guanylatecyclase-dependent). Although these vessel responses to LNO₂ suggest .NOas the mediator of guanylate cyclase activation, the identity of theproximal LNO₂-derived, cGMP-dependent signaling molecule was notdirectly identified (5).

Nitric oxide, synthesized by three different nitric oxide synthaseisoforms, was first shown to mediate endothelial-dependent relaxationvia reaction with the heme iron of guanylate cyclase and subsequentactivation of cGMP-dependent protein kinases (6). Subsequent to thisdiscovery, there has been a growing appreciation that the cell signalingactions of .NO are also transduced by secondary products derived fromredox reactions of .NO. These redox reactions yield a variety of oxidesof nitrogen displaying both unique and overlapping reactivities that canregulate differentiated cell function via both cGMP- andnon-cGMP-dependent mechanisms. These products include nitrite (NO₂ ⁻),.NO₂, peroxynitrite (ONOO⁻), nitrosothiols (RSNO) and dinitrogentrioxide (N₂O₃). These reactive species serve to transduce the cellsignaling actions of .NO by inducing changes in target moleculestructure and function via oxidation, nitration or nitrosation reactions(7,8).

The lipophilicity and intrinsic chemical reactivities of .NO facilitatemultiple interactions with lipids that impact both cellular redox and.NO signaling reactions. For example, .NO concentrates in membranes andlipoproteins, where it more readily reacts with oxygen to yieldoxidizing, nitrosating and nitrating species such as N₂O₃ and N₂O₄(9-11). In these lipophilic compartments, .NO can react with lipidperoxyl radicals (LOO.) at diffusion-limited rates, readilyout-competing tocopherols and ascorbate for the scavenging ofintermediates that would otherwise propagate lipid oxidation. In thisregard, .NO displays an oxidant-protective, anti-inflammatory role(12,13). Of relevance to inflammatory signaling, heme andnon-heme-containing peroxidases and oxygenases that catalyze physiologicand pathologic fatty acid oxygenation reactions also catalyticallyconsume .NO during enzyme turnover [e.g., lipoxygenases (14,15),cyclooxygenase (16) and myeloperoxidase (17)]. The reaction of .NO withthese enzymatic catalysts and free radical intermediates of fatty acidoxygenation in turn inhibits rates of fatty acid oxygenation productformation. The convergence of .NO and fatty acid oxygenation reactionsthus can influence the steady state concentration of both .NO andeicosanoids in a concerted fashion.

Redox reactions of .NO frequently induce the chemical modification oftarget molecules, including the nitrosylation (addition of .NO) of hemeproteins (18), the nitrosation (addition of the nitroso group NO) ofthiol substituents (7) and the nitration (addition of the nitro groupNO₂) of protein tyrosine residues and DNA bases (8). Herein, the presentinvention shows that LNO₂, a product of .NO-dependent unsaturated fattyacid nitration reactions that is abundant in red cells and plasma,decays in aqueous milieu to release .NO. This generation of .NO by LNO₂is inhibited by aprotic environments, a milieu that concomitantlystabilizes LNO₂. Moreover, it is shown that UV photolysis and I₃ ⁻-basedchemiluminescence approaches currently used to quantify .NO derived fromprotein heme-nitrosyl, RSNO and N-nitrosamine (RNNO) derivatives, alsofacilitate .NO release from LNO₂. This complicates the interpretation ofquantitative and qualitative results from the application of theseanalytical systems in biological preparations. In aggregate, theseresults reveal that nitroalkene derivatives of fatty acids serve totransduce the signaling actions of .NO via pathways subject toregulation by the relative distribution of LNO₂ to hydrophobic versusaqueous micro environments.

Materials—

Horse heart myoglobin, octyl-β-glucopyranoside (OG) andoctyl-thio-β-glucopyranoside (OTG), carboxy2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO),diethylenetriaminepentaacetate (DTPA), Na₂HPO₄ and sodium dithionitewere from Sigma (St Louis, Mo.). LNO₂ and [¹³C]LNO₂ were synthesized andpurified as previously described (1). Metmyoglobin was reduced usingsodium dithionite, desalted by exclusion chromatography on a SephadexPD-10 column and further oxygenated by equilibration with 100% oxygen.

Electron Paramagnetic Resonance—

EPR measurements were performed at room temperature using a BrukerElexsys E-500 spectrometer equipped with an ER049X microwave bridge andan AquaX liquid sample cell. The following instrument setting were used:modulation frequency, 100 kHz; modulation amplitude, 0.05 G; receivergain, 60 dB; time constant, 1.28 ms; sweep time, 5.24 s; center field,3510 G; sweep width, 100 G; power, 20 mW; scan parameter, 16 scans.

Spectrophotometry—

The UV spectrum of LNO₂ and repetitive scans of LNO₂ decay kinetics werecollected using a Hitachi UV 2401 PC spectrophotometer. Apparent .NOformation was calculated from extents of oxymyoglobin oxidation in thevisible wavelength range (spectrum) and at 580 nm (kinetic mode).Initial oxymyoglobin oxidation was calculated using a UV probe Version1.10 (ε₅₈₀ 14.4 mM⁻¹ cm⁻¹). Decomposition of the NO₂ group was followedat 268 nm and the appearance of oxidized products at 320 nm.

Liposome Preparation—

Reverse phase evaporation liposomes were formed fromdipalmitoylphosphatidylcholine (DPPC), cholesterol and stearylamine(4:2:1 mole ratio) following an established procedure (19). BrieflyDPPC, cholesterol and stearylamine were dissolved in CHCl₃ and sonicatedwith 10 mM KPi buffer (CHCl₃/KPi, 2:1, v/v). The organic solvent wasthen removed by evaporation under reduced pressure at 45° C. Theliposomes were allowed to anneal for 12 h at room temperature thencentrifuged and the pellet resuspended in the experimental buffer.

Chemiluminescence and UV Photolysis Analyses—

For direct detection of .NO release, LNO₂ (75 μM) was incubated directlyor with different additions (sulfanilamide, 1.5% w/v in 2 M HCl, 5 min,25° C., with or without HgCl₂, 50 mM) under aerobic conditions in acapped vial for 3 min. The gas phase was then injected into achemiluminescence detector (ANTEK Instruments, Houston, Tex.).Additionally, known concentrations of DEA-NONOate (in 10 mM NaOH) wereadded to a capped vial containing 0.5 M HCl. NOx concentration profilesof plasma samples were performed by .NO chemiluminescence analysis.Measurement of putative NO₂ ⁻, RSNO, and other .NO derivatives presentin plasma was performed by using an I₃ ⁻-based reducing system aspreviously (20,21). Rats were treated by intraperitoneal injection of 50mg/kg E. coli LPS and 5 hr later blood was collected in EDTAanticoagulation tubes following cardiac puncture. Following removal ofred cells by centrifugation (500 g×10 min), plasma samples werepretreated with either sulfanilamide (final concentration 1.5% w/v in 2M HCl, 5 min, 25° C.) with or without HgCl₂ (50 mM) prior to injectinginto the chemiluminescence detector to measure NO₂ ⁻ and HgCl₂-resistantNO_(x) derivatives, respectively. For UV photolysis studies, awater-cooled reaction chamber was filled with 1 ml of phosphate buffer(50 mM, pH 7.4 containing 10 μM DTPA) and continuously bubbled withargon. The chamber was illuminated using an ILC PS300-1A xenon arcsource (ILC Technology, Sunnyvale, Calif.). Samples were injected intothe reaction chamber through an air-tight septum and released .NO waspassed to the reaction chamber of a Sievers .NOA 280 NO analyzer anddetected by chemiluminescence after reaction with ozone (O₃).

Mass Spectrometric Analysis—

LNO₂ was extracted using the method of Bligh and Dyer (22). Duringextraction, [¹³C]LNO₂ was added as internal standard and the LNO₂content of samples quantified using LC/MS/MS (1). Qualitative andquantitative analysis of LNO₂ by ESI MS/MS was performed using a hybridtriple quadrupole-linear ion trap mass spectrometer (4000 Q trap,Applied Biosystems/MDS Sciex) as described (1). For the detection andcharacterization of L(OH)NO₂, the hydration product of LNO₂ generated bya Michael-like addition between H₂O and the nitroalkene, LNO₂ (3 μM) wasincubated at 25° C. for 60 min in 100 mM phosphate buffer containing 100μM DTPA pH 7.4 and extracted (Bligh and Dyer). L(OH)NO₂ was detectedusing a multiple reaction monitoring (MRM) scan mode by reportingmolecules that undergo an m/z 342/295 mass transition. This methodselects (m/z 342) in the first quadrupole, consistent with the precursorion, and following collision-induced dissociation (CID) yields in Q3 aspecies (m/z 295) consistent with loss of the nitro group ([M-(HNO₂)]⁻).Presence of the nitrohydroxy-adduct was confirmed by product ionanalysis of m/z 342. The degradation of LNO₂ to secondary products wasfollowed in negative ion mode after chloroform extraction and directinjection into an ion trap mass spectrometer with electrosprayionization (LCQ Deca, ThermoFinnigan).

Characterization of .NO Release from LNO₂—

cPTIO is a selective spin trap for .NO (k=10⁴ M⁻¹ s⁻¹, (23), with theproduct of this reaction, cPTI, displaying a characteristic EPRspectrum. In order to determine if .NO is derived from LNO₂, it wasincubated at 25° C. for different times in 100 mM phosphate buffercontaining 100 μM DTPA, pH 7.4 in the presence of cPTIO. This resultedin a time-dependent decrease of the characteristic five peak cPTIOsignal and the appearance of a new signal ascribed to cPTI (FIG. 20A).This release of .NO by LNO₂ was concentration-dependent and followedfirst order decay kinetics for LNO₂. Due to limitations of the .NO-cPTIOreaction for quantitating yields of .NO, oxymyoglobin was utilized tomeasure .NO release rates (24).

LNO₂ was incubated with oxymyoglobin in 100 mM phosphate buffercontaining 100 μM DTPA, pH 7.4 and .NO-dependent oxymyoglobin oxidationwas followed spectrophotometrically. LNO₂ oxidized oxymyoglobin in adose- and time-dependent fashion, yielding metmyoglobin as indicated bythe spectral changes depicted in FIG. 20B. The apparent rate constantfor .NO release by LNO₂, calculated from the oxidation of oxymyoglobinto metmyoglobin, was k=9.67×10⁻⁶ s⁻¹ (FIG. 20C). To monitor theconcomitant decomposition of the parent LNO₂ molecule, its UV spectrumwas first analyzed. LNO₂ displays a characteristic absorbance spectrumwith a peak at 268 nm, ascribed to the π electrons of the NO₂ group.During aqueous LNO₂ decay, this maximum decreases and a new maximumappears at 320 nm, corresponding to a mixture of oxygen and conjugateddiene-containing products not yet fully characterized by massspectroscopy (FIG. 20D). The decrease in absorbance at 268 nm paralleled.NO release, as detected by both EPR and oxymyoglobin oxidation.

Nitrite Formation During LNO₂ Decomposition—

During LNO₂-dependent .NO formation, measured via oxymyoglobin oxidationand MS analysis of LNO₂ parent molecule loss in aqueous buffers, thestable .NO oxidation product NO₂ ⁻ accumulates with time (FIG. 21). Therelease of .NO from LNO₂ was maximal at pH 7.4 (22), suggesting a rolefor protonation and deprotonation reactions in .NO formation from LNO₂.

Chemiluminescence Analysis of LNO₂-Derived .NO—

Gas phase O₃-mediated chemiluminescence detection of NO is a highlysensitive and specific method for detecting .NO. LNO₂ was incubated incapped vials in 100 mM phosphate buffer, 100 μM DTPA, pH 7.4 in air andthe gas phase directly injected into the detector. The .NO-dependentchemiluminescence yield was a function of concentration of DEA-NONOateand LNO₂ concentrations, studied separately (FIG. 23A).Chemiluminescence was also time-dependent, increasing with time of LNO₂decay prior to gas sampling from vials.

UV photolysis has been used to quantitate RSNO derivatives of proteinsand other NO-containing biomolecules (25,26). When LNO₂ (4 nmol) wassubjected to UV photolysis in concert with .NO chemiluminescencedetection, UV light exposure stimulated .NO release from LNO₂ (FIG.23B). The .NO chemiluminescence response to NO₂ ⁻ (4 nmol) added tosamples being subjected to UV photolysis and repetitive LNO₂ additionwas also examined to address the possibility that LNO₂-derived NO₂ ⁻formed during decay reactions might have accounted for some fraction ofnet chemiluminescent yield; it did not.

Appreciating that nitroalkene derivatives of red cell membrane andplasma fatty acids are present in human blood, whether LNO₂-derived .NOhas the potential to interfere with the chemiluminescent detection ofNO₂ ⁻, RSNO, RNNO or NO-heme compounds in plasma, when also analyzed viaa triiodide (I₃ ⁻)-based reaction system (21) was examined Plasma fromLPS-treated rats was used to exemplify this reaction system, since LPStreatment of rodents induces a robust elevation in plasma biomoleculeNO-adduct levels (27). First, plasma was directly injected into thedetector chamber and I₃ ⁻ reagent added, yielding a signal indicative ofnet plasma NO₂ ⁻, RSNO, RNNO and NO-heme compounds (FIG. 23C, peak 1).Then, plasma treated with acidic sulfanilamide (which removes NO₂ ⁻) wasinjected, giving a peak of lower intensity after I₃ ⁻ reagent addition,indicative of RSNO and putative RNNO derivatives (20,28). Finally, aplasma sample treated with acidic sulfanilamide and HgCl₂ was injected,which resulted in an even smaller peak following I₃ ⁻ reagent addition(FIG. 23C, peak 3). This latter peak has been referred to asHg-resistant RNNO derivatives (20,28). Using this strategy andcombination of reagents, LNO₂ pretreated with acidic sulfanilamide andHgCl₂ also generated .NO chemiluminescence for extended periods of timefollowing I₃ ⁻ addition (FIG. 23C, LNO₂ inset).

Hydrophobic Stabilization of LNO₂—

The observation that LNO₂ is stable in organic solvents such asn-octanol, undergoing decay only after solvation in aqueous solutions,led us to analyze rates of .NO formation from LNO₂ in the presence ofnon-ionic detergents. The formation of .NO was followed by EPR(measuring cPTI formation) in the presence of different concentrationsof octyl-β-glucopyranoside (OG) and octyl-thio-β-glucopyranoside (OTG).The rate of .NO release was constant and not influenced by thesedetergents until the critical micellar concentration (CMC) for each wasachieved, after which .NO formation was inhibited as the volume of thehydrophobic environment increased (FIG. 24A). Similar results wereobtained when measuring .NO formation via conversion of oxymyoglobin tometmyoglobin. Apparent .NO release rates remained constant until OGconcentration reached ˜2.8 mg/ml (CMC=2.77 mg/ml (11)). For OTG,inhibition of LNO₂-dependent .NO release occurred at ˜7 mg/ml (CMC=7.8mg/ml (11)) (FIG. 24B). To further confirm that LNO₂ was protected inlipophilic environments, LNO₂ decomposition was followed by UVabsorbance at 268 nm and 320 nm (FIGS. 24 C-D). The inhibition of theNO₂ group loss at 268 nm was paralleled by inhibition of the formationof oxidation products at 320 nm, similarly paralleling thedetergent-induced inhibition of .NO release observed by EPR andoxymyoglobin-based detection. The micellar stabilization of LNO₂ wasalso documented in OTG-containing buffers by MS-based quantification ofLNO₂ after Bligh and Dyer extraction (FIG. 25A). Assuming rapidpartitioning of LNO₂ between the aqueous and hydrophobic compartments,and that LNO₂ decay occurs only in the aqueous compartment, it can beshown that the rate of reaction v is given by the relationship:

$v = \frac{k\left\lbrack {L{NO}}_{2} \right\rbrack}{1 + {\alpha\left( {K - 1} \right)}}$where k is the rate constant for aqueous breakdown, α is the fraction oftotal volume that is the hydrophobic volume, and K (hydrophobic/aqueousconcentration ratio) is the partition constant for LNO₂. Thus, a plot of1/v vs. a will yield a linear plot with slope divided by y-axisintercept equal to K−1. FIG. 25B shows this plot for OG and OTG,yielding values for K of 1580 and 1320 respectively.

For evaluating the stability of LNO₂ in bilayers rather than micelles,phosphatidylcholine-cholesterol liposomes prepared by reverse phaseevaporation were utilized (3:1). This alternative hydrophobic bilayerenvironment also resulted in a dose-dependent inhibition of the releaseof .NO from LNO₂, as detected by EPR analysis of cPTI formation fromcPTIO (FIG. 25C).

The decay of LNO₂ in aqueous solutions results in the formation ofmultiple secondary fatty acid-derived products, as well as .NO. Onepathway that may be involved in aqueous LNO₂ decay is the Michael-likeaddition reaction with H₂O at the α carbon of the nitroalkene moiety. Totest this possibility, and the influence of micellar stabilization ofLNO₂, the formation of nitrohydroxylinoleic acid (L(OH)NO₂, m/z 342) wasanalyzed by MS. LNO₂-derived L(OH)NO₂ was evident after 60 minincubation in aqueous buffer at pH 7.4, with a concomitant decrease inLNO₂ levels (m/z 324). Addition of OTG at a concentration above the CMCsignificantly decreased extents of L(OH)NO₂ formation (FIG. 26A-C).Product ion analysis of L(OH)NO₂ generated a pattern of CID product ionsthat indicate the presence of two predominant regioisomers consistentwith the heterolytic scission products of L(OH)NO₂,9-hydroxy-10-nitro-12-octadecaenoic acid (m/z 171) and12-hydroxy-13-nitro-9-octadecaenoic acid (m/z 211, FIG. 26 D-E).

The release of .NO by LNO₂ via a modified Nef reaction mechanism wasfurther supported by detecting an aqueous degradation product with m/z293 (FIG. 27). This mass to charge ratio is consistent with theformation of a conjugated ketone (Scheme 2 (FIG. 29), Stage 2). Alsopresent in the mass spectrum is a peak for the vicinal nitrohydroxyadduct (m/z 342) and minor peaks corresponding to the hydroxy and peroxyderivatives of LNO₂ (m/z 340 and 356, respectively).

Nitrolinoleic acid is a pluripotent signaling molecule that exerts itsbioactivity by acting as a high affinity ligand for PPARγ (2),activating protein kinase signaling cascades and, as shown herein, byserving as a hydrophobically-stabilized reserve for .NO. The activationof PPARγ-dependent gene expression by LNO₂ requires this ligand to bestabilized and transported as the intact nitroalkene to the nuclearreceptor (2). The mechanism(s) involved in protein kinase activation byLNO₂ remain unclear, but can include direct ligation of receptors at theplasma membrane and/or covalent modification and activation of signalingmediators via Michael addition reactions. Current data reveals that thesignaling actions of LNO₂ are multifaceted, with the activation ofprotein kinases and/or PPAR receptor activation not fully explainingobserved cellular responses, such as the stimulation of cGMP-dependentvessel relaxation (5). The observation herein that LNO₂ decay yields .NOand that LNO₂ is subject to hydrophobic stabilization thus lendsadditional perspective to our understanding of how compartmentalizationwill influence the nature of cell signaling reactions mediated by fattyacid nitroalkene derivatives.

A central challenge in detecting .NO generation by relativelyslow-releasing compounds (e.g., RSNO and organonitrate derivatives) isthe risk of lack of specificity and sensitivity. This is especially thecase when concurrent oxygen, heme, lipid, protein and probe-relatedredox reactions are possible. Quantitative rigor is also always aconcern. To circumvent these problems, multiple approaches for thequalitative and quantitative detection of .NO generation by LNO₂ wereemployed herein. The release of .NO by LNO₂ was assessed quantitativelyby spectrophotometric analysis of oxymyoglobin oxidation. Additionalqualitative proof of LNO₂-derived .NO release came from EPR analysis of.NO-dependent cPTI formation, .NO-dependent chemiluminescence followingreaction with O₃ and mass spectroscopic detection of anticipated decayproducts of LNO₂. Also, in aqueous solutions and in the absence ofalternative reaction pathways, 4 mol .NO react with 1 mol O₂ toultimately yield 4 mol NO₂ ⁻. Thus, formation of NO₂ ⁻ was used asadditional evidence for .NO formation. The yield of NO₂ ⁻ during LNO₂decay was 3.5 fold lower than predicted from more direct .NOmeasurements based on oxymyoglobin oxidation. Several explanations canaccount for this apparent discrepancy. First, in the absence of .NOscavengers, .NO rapidly equilibrates with the gas phase, thus decreasing.NO available for oxidation to NO₂ ⁻. Second, .NO reactions withcarbonyl, hydroxyl and peroxyl radicals are extremely fast [k>˜1×10¹⁰M⁻¹ s⁻¹, (29)]. These free radical intermediates are likely formedduring LNO₂ decomposition, as evidenced by products with mass to chargeratio 340 and 356 (FIG. 27). Thus, products of the reaction of thesespecies with .NO may not contribute to NO₂ ⁻ formation. Overall,multiple independent criteria support the capacity of LNO₂ to release of.NO.

The gas-phase chemiluminescence reaction of .NO with O₃ is a highlysensitive and specific method for detecting .NO and nitroso-derivativesof biomolecules. One widely utilized analytical strategy relies on thereductive cleavage of NO₂ ⁻ and nitroso-derivatives by I₃ ⁻. Treatmentof samples with acidic sulfanilamide and HgCl₂ permits additionaldiscrimination between heme-NO, NO₂ ⁻, and putative RSNO and RNNOderivatives (20, 21, 25-28). The latter HgCl₂-resistant species[proposed as RNNO, (20)] may be best termed XNO_(x) at this juncture,since LNO₂ also yields O₃ chemiluminescence following reaction withacidified sulfanilamide and HgCl₂ prior to injecting intoiodine/triiodide mixtures and the detection chamber. These data revealthat a contribution of fatty acid nitroalkene derivatives to themeasurement of various tissue biomolecule NO derivatives mustadditionally be considered. Of additional interest, the UV photolysisapproach for NO_(x) detection in biological samples directly stimulatesdecay of LNO₂ to yield .NO. This new insight thus raises significantconcern about the accuracy of reported concentrations for .NO-derivedspecies using UV photolysis, since nitrated fatty acids are the mostprevalent bioactive oxides of nitrogen yet found in vivo (1). Proteinfractionation via solvent extraction (e.g., acetone) prior to analysisof .NO derivatives in biological samples does not eliminate thepossibility that nitrated fatty acids are a source of “detectable” orRSNO-like .NO formation by UV photolysis, as LNO₂ and other nitroalkenesreadily partition into the polar phase of many extraction strategiesincluding those employing acetone.

The observation that LNO₂ undergoes decay reactions to yield .NO inaqueous solution initially raised concern regarding how a significantand consistent LNO₂ content in plasma and red cells of healthy humanscould be detected at near-micromolar concentrations (1). Appreciatingthat synthetic LNO₂ is stable in methanol suggested that the ionicmicroenvironment in which LNO₂ was solvated would significantly modulatestability. To first address the possibility that LNO₂ is stabilized byhydrophobic environments reminiscent of membranes and lipoproteins, itwas observed that .NO release from LNO₂ was inhibited upon LNO₂solvation in n-octanol. Further analysis using non-ionic detergentmicelles, wherein the relatively hydrophobic NO₂ group of LNO₂ isexpected to partition into non-polar microenvironments, revealed thatLNO₂ decomposition and .NO release was inhibited (FIG. 24). Importantly,this occurred at and above the CMC of each detergent and lipid studied.Similar results were obtained using DPPC-phosphatidylcholine-cholesterol(3:1, mol/mol) liposomes, also revealing that LNO₂ is readilyincorporated into and stabilized by lipid bilayers (FIG. 25C). Thisstabilizing influence of liposomes, which have a very low CMC, occurredat low hydrophobic phase volumes. These data reveal that LNO₂ will bestable in hydrophobic environments and that cell membranes andlipoproteins can serve as an endogenous reserve for LNO₂ and itsdownstream cell signaling capabilities. Indeed, ˜80% of LNO₂ isesterified to complex lipids in blood, including phospholipids derivedfrom red cell membrane lipid bilayers (1). This further suggests thatduring inflammatory responses, esterases and A₂-type phospholipases mayhydrolyze and mobilize membrane-stabilized LNO₂ for mediating cellsignaling actions. This regulated disposition of LNO₂ in lipophilicversus aqueous environments thus represents a “hydrophobic switch” thatwill control the nature of LNO₂ signaling activity (Scheme 1 (FIG. 28)).

The mechanisms accounting for .NO release from organic nitrites andnitrates are controversial, appear to be multifaceted and remain to beincisively defined. For example, the nitrate ester derivativenitroglycerin (NTG) has been used as a vasodilator for more than acentury in the treatment of angina pectoris. Nitroglycerin does notdirectly decay to yield .NO or an .NO-like species that will activatesoluble guanylate cyclase (sGC), rather cellular metabolism is requiredto yield a species capable of .NO-like activation of sGC. While severalenzymes are identified as competent to mediate the denitration and“bioactivation” of NTG (e.g., xanthine oxidoreductase, cytochrome P450oxidase and reductase, old yellow protein and mitochondrial aldehydedehydrogenase-2), detailed insight is lacking as to unified redoxchemistry, enzymatic and cellular mechanisms accounting for a) the 3e-reduction of nitrate to an .NO-like species and b) the attenuated NTGmetabolism that occurs during nitrate tolerance (30).

The present report of non-enzymatic release of .NO from endogenous fattyacid nitroalkene derivatives (e.g., LNO₂) lends additional perspectiveto how nitric oxide synthase-dependent .NO signaling can be transduced.It is shown via 3 different analytical approaches that the product ofLNO₂ decay is unambiguously .NO. Mass spectrometric analysis and LNO₂decay studies reported herein, in concert with previous understanding ofthe chemical reactivity of nitroalkenes, reveals a viable mechanism forhow nitrated fatty acids can serve to transduce tissue .NO signalingcapacity (Schemes 1 and 2).

The release of .NO by a vicinal nitrohydroxy arachidonic acid derivativedetected in cardiac lipid extracts has been proposed (31). Thesederivatives induce vasorelaxation of rat aortic rings via possible.NO-dependent activation of guanylate cyclase. The intermediateformation of an analogous hydroxy derivative of nitrolinoleate,L(OH)NO₂, is documented herein to occur during LNO₂ decay in aqueousmilieu (FIG. 27). Fatty acid nitroalkene derivatives appear to beclinically abundant, since both nitro and nitrohydroxy derivatives ofall principal unsaturated fatty acids are present in healthy human bloodplasma and urine (32). Present results indicate that hydroxy derivativesof fatty acid nitroalkenes represent the accumulation of Michaeladdition-like reaction products with H₂O that are in equilibrium withthe parent nitroalkene and are not a direct precursor to .NO release.

A more viable mechanism accounting for .NO release by nitroalkenes issupported by 1) mass spectroscopic detection of expected decay productsand 2) the aqueous and pH dependency of this process (FIG. 22), withLNO₂ decay and consequent .NO release involving protonation anddeprotonation events. The mechanism accounting for .NO release bynitroalkenes is based on the Nef reaction (33,34), a standard reactionof organic nitro derivatives first described in 1894 (35).

The original Nef reaction entails complete deprotonation of an alkylnitro compound with base to yield the nitroanion, followed by quenchingwith aqueous acid to cause hydrolysis to the corresponding carbonylcompound and oxides of nitrogen. Most Nef reactions are now performedusing additional oxidants or reductants, rather than the simpleacid-base chemistry of the original reaction (36-44). There are a fewnoteworthy points about this proposed mechanism that relate to how .NOcan be ultimately produced. The nitrogen-containing product of theoriginal Nef reaction is N₂O, a stable oxide of nitrogen that would notbe a precursor to .NO under the neutral aqueous conditions used hereinto model biologically-relevant LNO₂ decay. The initial oxide of nitrogenformed, HNO, is unstable and quickly disproportionates to form N₂O asshown in Scheme 2 (FIG. 29) (Stage 2). While HNO (or the NO⁻ anion)might conceivably yield one electron and be oxidized to .NO, this is notexpected under neutral aqueous conditions. Alternatively, a nitrosointermediate formed during LNO₂ decay provides a plausible pathway toyield .NO. This nitroso intermediate is expected to have an especiallyweak C—N bond, easily forming .NO and a radical stabilized byconjugation with the alkene and stabilized by the OH group, a moietyknown to stabilize adjacent radicals.

In Scheme 2 (FIG. 29) (Stage 1) the vicinal nitrohydroxy fatty acidderivative is in equilibrium with the nitroalkene. This is possible fortwo reasons. First, the nitro group in the vicinal nitrohydroxy fattyacid makes the adjacent hydrogen very acid (pK_(a) ˜7-8), thusfacilitating formation of a significant amount of the nitronate anion atphysiological pH. The anion can then release hydroxide, which whenneutralized with the proton removed in the first step results in the netloss of neutral water. Second, the fatty acid nitroalkene is a strongelectrophile and can readily undergo Michael conjugate addition reactionwith the small amounts of hydroxide anion that are always present inaqueous solution under physiological pH conditions, explaining thefacile equilibrium of vicinal nitrohydroxy fatty acids with theircorresponding nitroalkene derivatives. In Scheme 2 (FIG. 29) (Stage 2),the lipid nitroalkene forms .NO as described above.

These proposed mechanisms for .NO formation from LNO₂ provided thetestable hypothesis for how nitrated fatty acids can serve as a sourceof .NO using simple acid/base chemistry with no additional oxidants orreductants. Mass spectrometric detection of expected oxidized fatty acidproducts and direct detection of .NO formation supported this pathway ofnitroalkene decay. This acid/base chemistry could also be employed byas-yet-undescribed enzymes that could catalyzephysiologically-significant extents of .NO release from the multiplelipid nitroalkene derivatives now being observed (32).

Therapeutic agents that release .NO are a rapidly expanding area of drugdesign. Dual-acting nitro and nitroso derivatives of existing drugs havebeen synthesized and are being studied for efficacy in treatingdiabetes, metabolic syndrome, hypertension and atherosclerosis. Theseinclude .NO-releasing statin derivatives and NO-non-steroidalanti-inflammatory derivatives such as NO-acetacylic acid, NO-ibuprofenand NO-piroxicam. These adducts were devised based on the precept thatan .NO donor moiety will augment therapeutic breadth and value. Thisclass of pharmaceuticals are of particular relevance when alterations inendogenous .NO signaling contributes to tissue pathogenesis. In thisregard, LNO₂ shares similarities with these classes of “chimeric”inflammatory-regulating compounds, as LNO₂ is a potent endogenous PPARγagonist that rivals extents of PPARγ activation induced by similarconcentrations of thiazolidinediones (2). Herein, the present inventionshows that LNO₂ also has the capability to release .NO in a regulatedmanner. Thus, the potential signaling actions of LNO₂ are expected to bepluripotent in nature.

In summary, .NO-mediated oxidative reactions with unsaturated fattyacids yield nitroalkene derivatives. Once formed, nitrated fatty acidsare hydrophobically stabilized by lipid bilayers and lipoproteins oralternatively, can be redistributed to aqueous environments to release.NO via a Nef-like reaction. In its native form, LNO₂ also activatesnuclear PPAR receptor-mediated regulation of gene expression. Thesecombined actions are expected to transduce the salutary inflammatorysignaling reactions that have been described for both .NO and LNO₂.Because LNO₂ production is increased by oxidative inflammatoryreactions, this species thus represents an adaptive mediator thatregulates potentially pathogenic tissue responses to inflammation.

Throughout Example 4, several publications have been referenced. Thesepublications are listed in the Reference List for Example 4. Thedisclosures of these publications in their entireties are herebyincorporated by reference into this application in order to more fullydescribe the compounds, compositions and methods described herein.

REFERENCE LIST FOR EXAMPLE 4

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T., Wang, X., Reiter, C. D., Yang, B. K., Vivas, E.    X., Bonaventura, C., and Schechter, A. N. (2002) J. Biol. Chem. 277,    27818-27828-   22. Bligh, E. G. and Dyer, W. L. (1959) Can. J. Biochem. Physiol.    37, 911-917-   23. Akaike, T., Yoshida, M., Miyamoto, Y., Sato, K., Kohno, M.,    Sasamoto, K., Miyazaki, K., Ueda, S., and Maeda, H. (1993)    Biochemistry 32, 827-832-   24. Hogg, N., Singh, R. J., Joseph, J., Neese, F., and    Kalyanaraman, B. (1995) Free Radic. Res. 22, 47-56-   25. McMahon, T. J., Moon, R. E., Luschinger, B. P., Carraway, M. S.,    Stone, A. E., Stolp, B. W., Gow, A. J., Pawloski, J. R., Watke, P.,    Singel, D. J., Piantadosi, C. A., and Stamler, J. S. (2002) Nat.    Med. 8, 711-717-   26. Stamler, J. S., Jaraki, O., Osborne, J., Simon, D. I., Keaney,    J., Vita, J., Singel, D., Valeri, C. R., and Loscalzo, J. (1992)    Proc. Natl. Acad. Sci. U.S.A. 89, 7674-7677-   27. Crawford, J. H., Chacko, B. K., Pruitt, H. M., Piknova, B.,    Hogg, N., and Patel, R. P. (2004) Blood 104, 1375-1382-   28. Janero, D. R., Bryan, N. S., Saijo, F., Dhawan, V., Schwalb, D.    J., Warren, M. C., and Feelisch, M. (2004) Proc. Natl. Acad. Sci.    U.S.A. 101, 16958-16963-   29. Padmaja, S, and Huie, R. E. (1993) Biochem. Biophys. Res.    Commun. 195, 539-544-   30. Thatcher, G. R., Nicolescu, A. C., Bennett, B. M., and    Toader, V. (2004) Free Radic. Biol. Med. 37, 1122-1143-   31. Balazy, M., Iesaki, T., Park, J. L., Jiang, H., Kaminski, P. M.,    and Wolin, M. S. (2001) J. Pharmacol. Exp. Ther. 299, 611-619-   32. Baker, P. R. S., Lin, Y., Schopfer, F. J., Woodcock, S. T.,    Long, M. H., Batthyany, C., Iles, K. E., Baker, L. M. S., Sweeney,    S., Braunchaud, B. P., Chen, Y. E., and Freeman, B. A. (2005) J Biol    Chem Submitted,-   33. Pinnick, H. W. (1990) Organic Reactions 38, 655-792-   34. Ballini, R. and Petrini, M. (2004) Tetrahedron 60, 1017-1047-   35. Nef, J. U. 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Example 5

The nitroalkenes of the present invention also display regulatoryactions towards the expression of genes related to inflammatoryresponses, cell growth, cell differentiation, cell signaling, cell death(apoptosis) and metabolism. For example, nitrolinoleate regulates geneexpression in human vascular endothelial cells and macrophages. Cellswere stimulated with vehicle and linoleic acid (2.5 μM, the latter twotreatments as controls) and LNO2 (2.5 mM) for 24 hours. The total RNAwas purified with a Qiagen Kit and analyzed with a Whole Human GenomeOligo Array (G4112A, 41K Genes, Agilent). Results showed thatnitrolinoleate regulates >5000 genes, with statistically-significantup-regulation of >2300 genes and down-regulation of >3000 genes. Incontrast, the native (precursor) fatty acid, linoleic acid, onlyregulates 14 genes, with an up-regulation of 5 genes and thedown-regulation of 9 genes.

Therefore, the present invention also provides for the selectiveregulation of clinically significant genes by nitrolinoleate. Table 5lists examples of genes down-regulated by nitrolineate that areassociated with cell signaling, growth, differentiation, metabolism,inflammatory responses, migration and apoptosis. Table 5 also includesgenes involved in the Jak/STAT signaling pathway, the NF-KB pathway andthe P13K/Akt pathway. The genes listed in Table 5 can be involved in oneor more processes associated with cell signaling, growth,differentiation, metabolism, inflammatory responses, migration andapoptosis. These genes can also be involved in one or more pathwaysselected from the group consisting of the Jak/STAT signaling pathway,the NF-KB pathway and the P13K/Akt pathway.

TABLE 5 Selected genes down-regulated by nitrolineate (>1.5 fold, p <0.05) RAC2; ras-related C3 botulinum toxin substrate 2 (rho 2.47 family,small GTP binding protein Rac2) CDC2; cell division cycle 2, G1 to S andG2 to M 2.75 CDC42; cell division cycle 42 (GTP binding protein, 1.65 25kDa) CCND3; Cyclin D3 1.51 CCNE1; Cyclin E1 1.58 CAMK1;calcium/calmodulin-dependent protein kinase I 2.24 TCF4; transcriptionfactor 4 2.63 PPARGC1B; peroxisome proliferative activated 2.20receptor, gamma, coactivator 1, beta RB1; retinoblastoma 1 (includingosteosarcoma) 2.06 TNF; tumor necrosis factor (TNF superfamily, member2) 2.01 TYK2; tyrosine kinase 2 2.00 STAT1; signal transducer andactivator of transcription 1.74 1, 91 kDa CAV3; caveolin 3 1.76 EDARADD;EDAR-associated death domain 1.69 DAPK2; death-associated protein kinase2 1.73 CASP8AP2; CASP8 associated protein 2 1.52 ITR; intimalthickness-related receptor 1.68

Table 6 lists additional genes that are also down-regulated bynitrolineate and are associated with cell signaling, growth,differentiation, metabolism, inflammatory responses, migration andapoptosis

TABLE 6 Down-regulated genes (>1.5 fold, p < 0.05). Genes associatedwith cell signaling, growth, differentiation, metabolism, inflammatoryresponses, migration and apoptosis TGFBI; 9.80 ILK; integrin-linkedkinase 2.94 MAPKAPK3; mitogen-activated 2.18 transforming proteinkinase-activated protein growth factor, kinase 3 beta-induced, 68 kDaPLAU; 7.58 EGR2; early growth response 2.65 ADRB2; adrenergic, beta-2-,2.04 plasminogen 2 (Krox-20 homolog, receptor, surface activator,Drosophila) urokinase VASP; 4.85 FDPS; farnesyl diphosphate 2.46 IL11;interleukin 11 2.02 vasodilator- synthase (farnesyl stimulatedpyrophosphate synthetase, phosphoprotein dimethylallyltranstransferase,geranyltranstransferase) AIF1; allograft 3.95 TGFB1; transforming growth2.46 PIK3CB; phosphoinositide-3- 1.93 inflammatory factor, beta 1(Camurati- kinase, catalytic, beta factor 1 Engelmann disease)polypeptide PTGS1; 3.95 IRS1; insulin receptor 2.40 TIMP2; tissueinhibitor of 1.92 prostaglandin- substrate 1 metalloproteinase 2endoperoxide synthase 1 (prostaglandin G/H synthase and cyclooxygenase)PTGES2; 1.59 PDGFRA; platelet-derived 2.36 TLR2; toll-like receptor 21.89 prostaglandin E growth factor receptor, alpha synthase 2polypeptide E2F2; E2F 3.70 RACGAP1; Rac GTPase 2.35 LDLR; low densitylipoprotein 1.77 transcription activating protein 1 receptor (familialfactor 2 hypercholesterolemia) NCOR2; nuclear 1.59 NCR1; naturalcytotoxicity 2.30 CARD9; caspase recruitment 3.33 receptor co-triggering receptor 1 domain family, member 9 repressor 2 SRF; serum1.56 PDGFA; platelet-derived 2.27 FLJ23091; putative NFkB 1.74 responsefactor growth factor alpha activating protein 373 (c-fos serumpolypeptide response element-binding transcription factor) BCL2A1; 3.50TIMP1; tissue inhibitor of 3.62 PDGFC; platelet derived growth 1.72BCL2-related metalloproteinase 1 factor C protein A1 (erythroidpotentiating activity, collagenase inhibitor) CARD9; 3.33 TIMP3; tissueinhibitor of 2.27 ARHGEF1; Rho guanine 1.64 caspase metalloproteinase 3(Sorsby nucleotide exchange factor recruitment fundus dystrophy, (GEF) 1domain family, pseudoinflammatory) member 9 FN1; fibronectin 1 3.33MMP19; matrix 1.58 JUNB; jun B proto-oncogene 1.60 metalloproteinase 19PPARD; peroxisome 3.12 CAMK1; 2.24 MYLK; myosin, light 3.13proliferative calcium/calmodulin- polypeptide kinase activated dependentprotein kinase I receptor, delta MAPKAPK3; mitogen- 2.18 ROCK1;Rho-associated, coiled- 1.59 activated protein kinase- coil containingprotein kinase 1 activated protein kinase 3 IGF1; insulin-like growthfactor 1.54 1 (somatomedin C) VEGFB; vascular endothelial 1.51 growthfactor B ECGF1; endothelial cell growth 1.73 factor 1 (platelet-derived)

Table 7 provides gene that are up-regulated by nitrolinoleate. Thesegenes are associated with apoptosis, cell signaling and/or growth.

TABLE 7 Genes up-regulated by nitrolinoleate. IL10RA; interleukin 10receptor, 3.173 alpha GADD45G; growth arrest and DNA- 3.155damage-inducible, gamma IRS2; insulin receptor substrate 2 3.243 HO-1,heme oxygenase-1 4.086 GADD45A; growth arrest and DNA- 2.823damage-inducible, alpha CASP1; caspase 1, apoptosis-related 2.572cysteine protease (interleukin 1, beta, convertase) CASP4; caspase 4,apoptosis-related 2.57 cysteine protease CCNA1; Cyclin A1 2.558 JUN;v-jun sarcoma virus 17 2.454 oncogene homolog (avian) CASP3; caspase 3,apoptosis-related 1.689 cysteine protease GADD45B; growth arrest andDNA- 1.674 damage-inducible, beta AATK; apoptosis-associated tyrosine1.503 kinase AMID; apoptosis-inducing factor 8.362 (AIF)-homologousmitochondrion- associated inducer of death

Example 6

The present invention also provides the activation of p-JNK and p-c-Junprotein kinases by nitrated lipids, by stimulating the phosphorylationof these kinases. The activation of these cell signaling mediators allowmodulation of cell signaling, growth, differentiation, metabolism,inflammatory responses, migration and apoptosis. FIG. 30 shows humanlung epithelial cell responses of p-JNK and p-c-JUN upon administrationof nitrolineate. The nitroalkenes of the present invention also activatethe ERK MAPK pathway in human lung epithelial cells, as shown by adramatic increase in ERK phosphorylation (e.g. activation) and thephosphorylation of its downstream target signaling protein, pELK,illustrated in FIG. 31. The nitroalkenes of the present invention alsoinhibit activity of NF-KB pathway as indicated by a) luciferase-linkedNFkB-response element reporter assay in response to the inflammatorymediator TNFα and b) direct analysis of the degradation f the NFkBinhibitor protein, IkB in response to the inflammatory mediator E. coliLPS (FIG. 32).

Example 7 Synthesis and Characterization of Nitro/Hydroxy Fatty Acids

The Michael addition reaction of water to nitroalkenes results information of nitro-hydroxy derivative (18:1, 18:2 and 18:3 nitro-hydroxyspecies shown in FIG. 33). These hydroxylated species are prepared byplacing nitroalkenes in basic aqueous conditions, isolation by solventextraction and purification by HPLC or thin layer chromatography. Fattyacid nitro-hydroxy derivatives can (a) display unique cell signalingactivities, (b) represent a more stable “storage form” of the PPARreceptor-avid nitroalkene parent molecule and (c) permit determinationof specific positional isomers of nitroalkenes by directing, uponhydroxylation, nucleophilic heterolytic scission between the nitro andhydroxy-bonded fatty acid alkene. Nitro isomer-specific fragments canthen be detected by mass spectrometry (FIG. 33).

Example 8 Addition of Glutathione to Nitrated Fatty Acids

In a neutral buffered solution, the thiol and nitrated FA (e.g.,nitrated oleate or linoleate) can be combined from equimolar to 10:1ratios of nitroalkene to thiol. Mass spectrometry shows nitrolinoleateforms a covalent Michael addition reaction product with the tripeptideglutathione (m/z 631.3, LNO2-GSH) (FIG. 34). This adduct can be“fingerprinted” by its source fragmentation in the mass spectrometer tothe precursors glutathione (m/z 306.3) and nitrolinoleate (m/z 324.2,LNO₂).

Throughout this application, various publications are referenced. Thedisclosures of these publications in their entireties are herebyincorporated by reference into this application in order to more fullydescribe the compounds, compositions and methods described herein.

Various modifications and variations can be made to the compounds,compositions and methods described herein. Other aspects of thecompounds, compositions and methods described herein will be apparentfrom consideration of the specification and practice of the compounds,compositions and methods disclosed herein. It is intended that thespecification and examples be considered as exemplary.

What is claimed:
 1. A method for producing a nitrated lipid, comprising(a) reacting an unsaturated lipid with a mercuric salt, a seleniumcompound, and a nitrating compound to produce a first intermediate, and(b) reacting the first intermediate with an oxidant.
 2. The method ofclaim 1, wherein step (a) is performed under anaerobic conditions. 3.The method of claim 1, wherein step (a) is conducted under anhydrousconditions.
 4. The method of claim 1, wherein the mercuric saltcomprises HgCl₂, Hg(NO₃)₂, or Hg(OAc)₂.
 5. The method of claim 1,wherein the selenium compound comprises PhSeBr, PhSeCl, PhSeO₂CCF₃,PhSeO₂H, or PhSeCN.
 6. The method of claim 1, wherein the nitratingcompound comprises a nitrite salt.
 7. The method of claim 1, wherein thenitrating compound comprises NaNO₂ or AgNO₂.
 8. The method of claim 1,wherein the oxidant comprises H₂O₂ or an organic hydroperoxide.
 9. Themethod of claim 1, wherein the lipid comprises 14:1, 16:1, 18:1,18:2,18:3, 20:4, or 22:6, the mercuric salt comprises HgCl₂, the seleniumcompound comprises PhSeBr, the nitrating compound comprises NaNO₂, andthe oxidant comprises H₂O₂.
 10. The method of claim 9, wherein step (a)is performed under anaerobic and anhydrous conditions.
 11. The method ofclaim 1, wherein when after step (b) two or more isomers of the nitratedlipid are produced, separating each isomer so that each isomer issubstantially pure.
 12. The method of claim 11, wherein the separationstep comprises chromatographing the isomers.